The Pennsylvania State University

The Graduate School

Eberly College of Science

MECHANISTIC ANALYSIS OF THE TRYPTOPHAN BIOSYNTHETIC

ENZYME INDOLE-3-GLYCEROL PHOSPHATE SYNTHASE

A Dissertation in

Chemistry

by

Margot J. Zaccardi

©2013 Margot J. Zaccardi

Submitted in Partial Fullfillment

of the Requirements

for the Degree of

Doctor of Philosophy

December 2013

 

 

The dissertation of Margot J. Zaccardi was reviewed and approved* by the following:

David D. Boehr

Assistant Professor of Chemistry

Dissertation Advisor

Chair of Committee

Scott S. Showalter

Assistant Professor of Chemistry

Phillip C. Bevilacqua

Professor of Chemistry

Craig Cameron

Professor of Biochemistry and Molecular Biology

Eberly Chair in Biochemistry and Molecular Biology

Barbara J. Garrison

Shapiro Professor of Chemistry

Head of the Chemistry Department

*Signatures are on file in the Graduate School

 

 

iii  

ABSTRACT

The design and production of enzymes that are capable of performing reactions in

an industrial setting is of profound importance for the advancement of technologies

including pharmaceuticals, biotechnology, agriculture, and materials. While some natural

enzymes have industrial relevance, the design of novel enzymes whose catalytic reactions

are not found in natural systems would greatly enhance the scope and efficiency of

industrial processes. The difficulty of engineering such enzymes is that it requires in

depth knowledge of all factors of catalysis and how they affect the reaction, including an

understanding of the natural enzyme scaffold to be used in new design. Indole-3-glycerol

phosphate synthase (IGPS), a tryptophan biosynthetic enzyme that catalyzes the ring

closure of 1-(o-carboxyphenylamino)-1-deoxyribulose 5-phosphate (CdRP) to form

indole-3-glycerol phosphate (IGP), is widely used as a scaffold in enzyme engineering

studies. However, the rate enhancements of these engineered enzymes are much lower

than those of natural enzymes. This work has analyzed the IGPS enzyme from the

thermophile Sulfolobus sulfataricus (ssIGPS) and gained a more complete understanding

of the kinetic and chemical mechanism for this enzyme that can be leveraged towards

enzyme engineering applications.

Steady-state kinetic assays were used to analyze wild type (WT) ssIGPS and

variants. The results showed a temperature dependent change in the rate-determining step

of the reaction; the ring closure is rate-determining at high temperatures and product

release is rate-determining at low temperatures. These studies also showed that

thermophilic ssIGPS and mesophilic IGPS from Escherichia coli (ecIGPS) have different

rate-determining steps at their biologically relevant temperatures.

 

 

iv  

In addition to examining the temperature dependence of ssIGPS, amino acid

substituted variants of ssIGPS were used to probe the chemical mechanism of the

enzyme. While a proposed mechanism had been previously published, this analysis

provides additional and important details that were formerly unknown. The general acid

and base in the dehydration step of the reaction were reassigned to Lys53 and Glu51, as

opposed to the previously assigned Lys110 and Glu159 (or Glu210). This assignment

allowed for a more complete view of catalysis. First, Lys110 initiates the reaction by

donating a proton to the C2’ carbonyl of the substrate, which allows the ring closure to

occur and form the fleetingly stable I1 intermediate, which then undergoes

decarboxlation to form the I2 intermediate. The I2 undergoes a reorientation in the active

site to properly align it for dehydration assisted by the general acid and base, Lys53 and

Glu51. This step renders the product and leaves Lys53 neutral allowing for efficient

product release.

Finally, the role of the active site loop residues in ssIGPS catalysis was examined.

Lys53, the general acid in the dehydration step, is located on the dynamic β1α1 loop, and

Phe89, important for substrate binding, is on the β2α2 loop. Arg54 and Asn90, also on

these loops, were found to be coevolving by statistical coupling analysis (SCA), and

molecular dynamics (MD) simulations predicted their motions were also correlated. To

further assess the role of these residues, kinetic analysis of amino acid substituted

variants was performed. The results show that the interaction between Arg54 and Asn90

is important for the dehydration step of the reaction, mainly in the correct function of the

general acid and base, Lys53 and Glu51. The results also suggest that these residues play

a role in conformational exchange, as the effect on catalysis is temperature dependent,

 

 

v  

suggesting that thermal energy at higher temperatures can help to overcome their

detrimental effect on catalysis. Together, these results provide a more complete

understanding of ssIGPS catalysis. The results can be leveraged towards the design of

novel enzymes as well as in the development of new antimicrobials.

 

 

vi  

TABLE OF CONTENTS

LIST OF FIGURES……………………………………………………………………....ix

LIST OF TABLES……………………………………………………………………….xii

LIST OF ABBREVIATIONS…………………………………………………………...xiv

ACKNOWLEDGEMENTS……………………………………………………………..xvi

Chapter 1 Introduction to Indole-3-glycerol Phosphate Synthase………………..………1

1.1 Progress Towards Engineering Enzymes with New Functions………………1 1.2 The Conserved (β/α)8-Barrel Protein Fold as an Enzyme Engineering

Scaffold……….…………………………………………………………..5 1.3 (β/α)8-Barrel Enzymes in Tryptophan Biosynthesis………………………….6 1.4 The Tryptophan Biosynthetic Enzyme Indole-3-glycerol Phosphate

Synthase…………………………………………………………………..8 1.5 Previous Knowledge on the IGPS Mechanism....……………………………11 1.6 Conclusions...………………………………………………………...............20 1.7 References ……………………………………………………………………22 Chapter 2 The Temperature Dependent Kinetic Mechanism of Thermophilic and

Mesophilic IGPS Enzymes……………………………………………………....30 2.1 Abstract............................................................................................................30 2.2 Introduction…………………………………………………………………..31 2.3 Experimental Methods.....................................................................................34 2.3.1 Cloning of ssIGPS and ecIGPS…………………………………….34 2.3.2 Overexpression and Purification of ssIGPS, ecIGPS,

and tmIGPS………………………………………………………35 2.3.3 Steady-state Kinetic Assays for IGPS……………...……………....37

2.3.4 Solvent Viscosity Effects, Solvent Deuterium Kinetic Isotope Effects, and pH Effects………………………………………..…………39 2.3.5 Synthesis of CdRP…………………........………………………....41 2.3.6 Circular Dichroism...........................................................................42

2.4 Results……………………………………………………………………….43 2.4.1 Steady-state Kinetics of ssIGPS…………………………………...43 2.4.2 Solvent Viscosity Effects, Solvent Deuterium Kinetic Isotope

Effects, and pH Effects..…………………………………………45 2.5 Discussion……………………………………………………………………54 2.5.1 Temperature Dependent Kinetic Mechanism of ssIGPS…………..54 2.5.2 Differences in the Rate-Determining Step of Thermophilic ssIGPS

and Mesophilic ecIGPS at their Adaptive Temperatures………...55 2.6 Conclusions…………………………………………………………………..56

 

 

vii  

2.7 References……………………………………………………………………56 Chapter 3 Functional Identification of the General Acid and Base in the Dehydration Step

of Indole-3-glycerol Phosphate Synthase Catalysis……………………………...59 3.1 Abstract............................................................................................................59 3.2 Introduction…………………………………………………………………..60 3.3 Experimental Methods….……………………………………………………63

3.3.1 Overexpression, Purification, and Kinetic Analysis of WT and Amino Acid Substituted IGPS.…………………………………..63

3.3.2 Overexpression and Purification of ε-13C-Lys Labeled ssIGPS…...64 3.3.3 Preparation of rCdRP………………………………………………65 3.3.4 13C-TROSY-HSQC Experiments on ssIGPS………………………66

3.4 Results………………………………………………………………………..67 3.4.1 Determination of the Rate-Determining Step of ssIGPS

Catalysis........................................................................................67 3.4.2 Analysis of Lys53 Indicates its Role as a General Acid…………...67 3.4.3 13C-Lys NMR to Determine pKas for Lys Residues in IGPS……...73 3.4.4 Analysis of Glu51 Identifies its Role as the General Base….……..81

3.5 Discussion……………………………………………………………………82 3.6 Conclusions…………………………………………………………………..89 3.7 References……………………………………………………………………89 Chapter 4 The Role of Active Site Loops in Catalysis by IGPS…..…………………….92

4.1 Abstract............................................................................................................92 4.2 Introduction…………………………………………………………………..92 4.3 Experimental Methods……………………………………………………….95 4.4 Results………………………………………………………………………..96 4.4.1 Investigation of Phe89 on the β2α2 Loop Identifies its Role in IGPS

Catalysis ………………………………...……………………….96 4.4.2 Interaction Between β1α1 and β2α2 Loops through Arg54 and

Asn90 is Important for Catalysis………………………………...98 4.4.3Analysis of Arg54Lys and Asn90Gln variants of ssIGPS………...103 4.4.4 Examination of the Interaction Between Coevolving Residues on the β2α2 Loop…………………………………………………..104 4.4.5 Structure and Stability of Loop Mutants………………………….105 4.5 Discussion…………………………………………………………………..107 4.6 Conclusions…………………………………………………………………110 4.7 References………………………………………………………………......111

Chapter 5 Conclusions………………………………………………………………….113

5.1 A New Understanding of Catalysis by IGPS……………………………….113 5.2 Implications for Understanding the Evolution of Thermophilic Versus Mesophilic Enzymes……………………………………………………115

 

 

viii  

5.3 Engineering New Indole Derivatives and Improving Industrial Indole Synthesis with Biocatalysts…………………………………………….116 5.4 Improving Novel Enzyme Engineering Efforts…………………………….117 5.5 Future Studies..……………………………………………………………..120 5.6 Conclusions…………………………………………………………………123 5.7 References…………………………………………………………………..123

Appendix Solvent Deuterium Kinetic Isotope Effect Analysis ………………..............126

 

 

ix  

LIST OF FIGURES

Figure 1.1: The three consecutive (β/α)8-barrel enzymes in tryptophan biosythnesis catalyze the fourth, fifth, and sixth steps of the pathway. PRAI (PDB 1Pll) catalyzes the amadori rearrangement of PRA to form CdRP. CdRP is then converted to IGP by IGPS (PDB 1Pll). Then, αTS (PDB 1V7Y) catalyzes the cleavage of IGP to form glyceraldehyde-3-phosphate and indole. ...........……...…………………………………..7 Figure 1.2: Indole-3-glyerol phosphate synthase (IGPS). (a) ssIGPS (PDB: 1IGP) is a (β/α)8-barrel enzyme that contains an additional 45 residue N-terminal extension. (b) IGPS catalyzes the conversion of CdRP to form IGP. The proposed mechanism contains three steps and two intermediates and utilizes a general acid and base (proposed as Lys110 and Glu159)……………………………………………………………………..12 Figure 1.3: Multiple sequence alignments between IGPS from S. sulfataricus (ssIGPS), Thermatoga maritima (tmIGPS), E. coli (ecIGPS), and Mycobacterium tuberculosis (mtIGPS). Secondary structure is denoted by boxes above the sequence with α-helices in green and β-sheets in blue. Conserved residues are bold and in blue. Catalytically relevant residues are denoted with a star. Sequence alignment was performed with Clustal Omega provided by The European Bioinformatics Institute at The European Molecular Biology Laboratory……………………………….................................………………..14 Figure 1.4: Active site of IGPS with reduced CdRP bound. Conserved and catalytically relevant residues are shown. Lys110 is the proposed general acid in the condensation and dehydration steps. Glu159 and Glu210 have both been proposed to act as general base. Phe89 and Arg182 are proposed to aid in substrate binding. Lys53 is also involved in substrate binding and may have additional roles in the chemistry. The role of Glu51 has not been extensively studied……………………………………………………………..15 Figure 1.5: IGPS in complex with rCdRP (PDB:1LBF) (yellow) and IGP (1A53) (blue) showing residues that interact with the anthranilate moiety. When CdRP binds in the active site, Trp8, Pro57, Phe89, Arg182, and Leu184 interact with the anthranilate. Conversely, when IGP binds, Phe89, Lys110, Phe112, Ile133, and Arg182 interact.…..18 Figure 1.6: Numbering for CdRP. Lys53 interacts with the C1 carboxyl and C3’ hydroxyl groups and is thought to aid in ring closure between C1 and C2’.....................19 Figure 2.1: Conserved structure and function of IGPS from E. coli (green) (PDB 1P11) and S. sulfataricus (blue) (PDB 1IGPS). Despite only 30% sequence identity and large differences in stability, ssIGPS and ecIGPS show strong structural similarity………….33 Figure 2.2: Standard curve of fluorescence units per nanomolar for converting cps/s to nM/s. The slope of the line (4036 cps/nM) was used to convert data for ssIGPS to the appropriate units. Curve was attained using IGPS from T. maritima, which does not display product inhibition..................................................................................................38

 

 

x  

Figure 2.3: Representative data for ssIGPS assays. (a) Progress curves for ssIGPS at 75 °C at varying concentrations of CdRP (100, 400, 800, 1000, and 2000 nM). (b) Michaelis-Menton curve for ssIGPS at 75 °C...................................................................40 Figure 2.4: The rate-determining step of the IGPS catalyzed reaction can be determined using SVE and SDKIE experiments. Substrate binding and product release (green) are viscosity sensitive and isotope insensitive. Ring closure (blue) is viscosity insensitive and isotope sensitive. The decarboxylation and dehydration (red) are viscosity and isotope insensitive………………………………………………………………………………..46 Figure 2.5: Solvent viscosity effects for ssIGPS. At 25 °C (blue) there is an SVE of 1.0 ± 0.2, wherease at 75 °C (black) the SVE is no longer present (-0.2 ± 0.1). The SVE is defined by the slop of the line for vo/vi versus ni/no. The results indicate that at 25 °C product release is rate-determining but as temperature increases to 75 °C product release is no longer rate-determining, and a chemical step becomes rate-determining.................47 Figure 2.6: The pH dependence of WT ssIGPS at (a) 37 °C (pKa1 7.5 ± 0.2, pKa2 8.8 ± 0.3) and (b) 75 °C (pKa1 5.6 ± 0.2, pKa2 8.7 ± 0.2) and (c) ecIGPS at 37 °C (pKa1 6.7 ± 0.1, pKa2 8.8 ± 0.1) show an ascending and descending limb consistent with general base and general acid involvement, respectively....…………………………………………...49   Figure 2.7: The rate-determining step for ssIGPS at higher temperatures involves a single proton transfer event. (a) The maximum catalytic turnover of ki/ko versus mole fraction D2O:H2O at both 37 °C (blue) and 75 °C (green) show a linear fit. (b) The square root of ki/ko versus mole fraction D2O:H2O at 37 °C and 75 °C show a quadratic fit. These results indicate that one proton transfer event is involved in the rate-determining step of the reaction, namely the proton transfer from the general acid in the condensation step of the reaction.............................................................................................................53   Figure 3.1: Accepted mechanism for IGPS suggests that the reaction proceeds in three steps: condensation, decarboxylation, and dehydration, with two distinct intermediates……………………………………………………………………………..61 Figure 3.2: The pH profiles suggest that Lys53 and Glu51 act as the general acid and base, respectively, in the dehydration step of IGPS catalysis. Shown are the pH curves for IGPS (a) WT (pKa1 5.6 ± 0.2, pKa2 8.7 ± 0.1), (b) Lys53Arg (pKa1 6.9 ± 0.1, pKa2 > 9), and (c) Glu51Gln (pKa2 6.51 ± 0.3)……………………………………………………..71 Figure 3.3: 1H-13C HSQC on ε-13CH2-Lys labeled ssIGPS shows eighteen resonances, which is consistent with the number of lysine residues in the enzyme………………….74 Figure 3.4: Overlay of 1H-13C-HSQC Spectra for ssIGPS at pH 7.0 (black) and pH 10.5 (red). At high pH, the changes in the spectrum are likely caused by denaturation of ssIGPS................................................................................................................................75

 

 

xi  

Figure 3.5: A representative plot of chemical shift versus pH for peak #2. The pKa value associated with this curve is 11.45 ± 0.14, although the change in chemical shift likely reflects denaturation of the enzyme rather than deprotonation of the lysine.....................76 Figure 3.6: pH dependence of the ssIGPS catalyzed reaction performed in H2O (pKa1 5.7 ± 0.1, pKa2 8.7 ± 0.1), shown in blue, and D2O (pKa1 5.3 ± 0.2, pKa2 8.9 ± 0.2), shown in green, display little difference in pKa values. Therefore, the use of D2O in NMR experiments does not explain the discrepancy in pKa values between the two methods…………………………………………………………………………………..78 Figure 3.7: Overlay of 1H-13C HSQCs of Lys53Arg (red) and WT ssIGPS (black) labeled with ε-CH2-Lys. Due to the low resolution of Lys53Arg ssIGPS spectrum, and its the poor alignment to WT, Lys53 and other resonances remain unassigned………………..79 Figure 3.8: The assigned role for the conserved and charged residues in the active site of IGPS. The ring closure is catalyzed by the general acid, Lys110, and assisted by Glu159 (blue). The dehydration is catalyzed be the general acid, Lys53 and general base, Glu51 (yellow). Arg182 and Glu210 are involved in substrate binding (orange)………………83 Figure 3.9: The modified mechanism of ssIGPS catalysis utilized Lys53 and Glu51 as the general acid and base pair in the dehydration step of the reaction. Additionally, the general base now attacks the amide hydrogen rather than the previously suggested alkyl hydrogen............................................................................................................................85 Figure 3.10: Rotation about the C3’-C4’ bond of ribose chain is required for the dehydration step in IGPS catalysis. Crystal structure of ssIGPS:IGP complex (Top) (PDB: 1LBF) shows the ligand bound in the active site such that Lys53 and Glu51 are not properly positioned for catalysis. The ribose chain must rotate (Bottom) to reposition the intermediate in the second binding pocket and allow for dehydration to form IGP…….86 Figure 3.11: Surface rendering of the IGPS binding pocket shows two distinct active sites for catalysis. In the first site (blue), Lys110 and Glu159 catalyze the ring closure step deep within the pocket. The intermediate then transitions to the second site (yellow), which is closer to where product exits the binding pocket, where Lys53 and Glu51 catalyze the dehydration step. (PDB: 1A53)…………………………………………….88

 

 

xii  

Figure 4.1: The catalytically important residues in the dehydration step of IGPS are found on the β1α1 and β2α2 loops, which interact through a hydrogen bond between Arg54 and Asn90. (a) The ssIGPS catalyzed reaction has two distinct binding pockets for the two reaction steps. In step one, Lys110 (cyan) initiates the ring closure and decarboxylation. Following the formation of the intermediate, the anthranilate moiety is transferred to the second site where Lys53 and Glu51 (yellow) act as the active site acid and base. The role of the β1α1 and β2α2 loops (blue) including the interaction between Arg54 and Asn90 is examined herein. (b) This interaction is thought to have functional significance in IGPS since it is coevolving amongst IGPS species and exhibits correlated motion in MD simulations. This interaction is in close proximity to the conserved residues Lys53 and Phe89………………………………………………………………..94 Figure 4.2: pH profiles of WT and Asn90Ala ssIGPS show changes to the activity of the general acid and base. pH profile for (a) WT (pKa1 5.6, pKa2 8.7) shows two pKa values associated with general acid/base catalysis, whereas (b) Asn90Ala (pKa1 7.26 ) show a loss in the second ionization that was previously attributed to Lys53. This finding indicates that the Asn90 variant is affecting the dehydration step of the reaction, and interfering with proper function of the general acid Lys53…………..………………...100 Figure 4.3: Asn90Ala affects the thermal stability but not proper folding for ssIGPS. (a) Thermal inactivation curves show an increase in the thermal stability of the Asn90Ala variant compared to WT. (b) Circular dichroism curves indicate that the changes in activity for the Asn90Ala variant are not caused by gross structural changes to the enzyme and can be attributed to changes in the reaction……………………………….106

 

 

xiii  

LIST OF TABLES

Table 1.1: Conserved active site residues in ssIGPS that are of interest to these studies

are shown along with their proposed roles in enzyme activity..........................................16

Table 2.1: Steady-state kinetic parameters for ssIGPS and ecIGPS at pH 7.5 indicate that

the rate-determining step changes as a function of temperature. ……......………………44

Table 2.2: pKa values for ssIGPS and ecIGPS..................................................................50

Table 3.1: Steady-state kinetics (at 75 °C) demonstrates that the dehydration step of

IGPS catalysis occurs through the general acid and base Lys53 and Glu51,

respectively. .........................................................………………...............……………..68

Table 3.2: pKa values for WT ssIGPS and Lys53Arg and Glu51Gln variants identify

Lys53 and Glu51 as the general acid and base in ssIGPS catalysis...................................72

Table 3.3: pKa values determined by 1H-13C HSQC on ε-13CH2-Lys labeled ssIGPS are

not in agreement with the pKa for the pH dependence of the enzymatic reaction

determined by steady-state kinetics……………………………………………………...77

Table 4.1: Steady-state kinetic parameters of WT ssIGPS and loop variants indicate an

important role for the β1α1 and β2α2 loop interaction in catalysis……………………...97

Table 4.2: pKa values for WT, Arg54Ala, and Asn90Ala indicate that the loop

interaction is important for general acid/base catalysis...................................................101

 

 

xiv  

LIST OF ABBREVIATIONS

Amp Ampicillin

BICINE N,N-Bis(2-hydroxyethyl)glycine

CD Circular dichroism

CdRP 1-o-carboxylphenylamino deoxyribulose 5-phosphate

CHES N-cyclohexyl-2-aminoethanesulfonic acid

CPS Counts per second

DTT Dithiothreitol

ecIGPS Indole-3-glycerol phosphate synthase from Escherichia coli

EDTA Ethylenediaminetetraacetic acid

EI Enzyme-intermediate complex

EP Enzyme-product complex

ES Enzyme-substrate complex

Hepes 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid

Hepps 3-[4-(2-hydroxyethyl)-1-piperazinyl] propanesulfonic acid

HPLC High pressure liquid chromatography

HSQC Heteronuclear single quantum coherence

I1 Intermediate 1 from the IGPS reaction

I2 Intermediate 2 from the IGPS reaction

IGP Indole-3-glycerol phosphate

IGPS Indole-3-glycerol phosphate synthase

IPTG Isopropyl β-D-1-thiogalactopyranoside

Kan Kanamycin

kcat Maximum catalytic turnover constant

KM Michaelis constant

LB Luria-Bertani

MD Molecular dynamics

MES 2-(N-morpholino)ethanesulfonic acid

mtIGPS Indole-3-glycerol phosphate synthase from Mycobacterium tuberculosis

NaBH4 Sodium borohydride

 

 

xv  

NAC Near attack conformer

NMR Nuclear magnetic resonance

P Product

PCR Polymerase chain reaction

PDB Protein data bank

PMSF Phenylmethanesulfonyl fluoride

PRA Phosphoribosyl anthranilate

PRAI n-Phosphoribosyl anthranilate isomerase

rCdrP Reduced CdRP

S Substrate

SCA Statistical coupling analysis

SDKIE Solvent deuterium kinetic isotope effect

SDS-PAGE Sodium dodecylsulfate-polyacrylamide gel electrophoresis

ssIGPS Indole-3-glycerol phosphate synthase from Sulfolobus sulfataricus

SVE Solvent viscosity effect

TB Tuberculosis

TIM Triose Phosphate Isomerase

tmIGPS Indole-3-glycerol phosphate synthase from Thermotoga Maritima

tr-NOE Exchange transferred-nuclear Overhauser effect

TrpC IGPS encoding gene

WT Wild type

αTS Alpha subunit of tryptophan synthase

 

 

xvi  

ACKNOWLEDGEMENTS

There are many people who deserve recognition for their support throughout my

graduate studies. I would first like to thank my advisor, David Boehr, whose mentorship

allowed me to develop into a thoughtful and independent scientist, as well as my

committee, Scott Showalter, Phil Bevilacqua, and Craig Cameron, for their thoughtful

comments on this work. I must also thank the other members of the Boehr laboratory,

especially Alyson Boehr, Jennifer Axe, Yan Mei Chan, Olga Manweiler, Laura Loggia,

and Alexander Chasin. Life in the Boehr lab as been an interesting and memorable

experience, and their support and entertainment has helped make the days more

interesting. I am honored to be the first person to complete my Ph.D. from the laboratory.

Without the use of the fluorometer in the Benkovic laboratory, I would not have been

able to complete this thesis, and so I would also like to thank Stephen Benkovic, Michelle

Spiering, and the other members of the Benkovic laboratory for allowing me into their

laboratory and for the generous loan of their instrumentation.

The mentorship I received at the Florida Institute of Technology from my

undergraduate research advisor, Dr. Joel Olson, is the reason I attended graduate school,

and also part of the reason I was able to succeed. He was the first person to spark my

curiosity for chemical research, and his famous words of wisdom, “research is like a

funnel,” have gotten me through the slow and tedious parts of this process and helped me

remember the bigger picture.

While the words presented in this dissertation reflect my scientific work, my

persistence in completing this thesis is largely due to those hours between experiments

spent with some amazing friends, especially Dr. Joy Gallagher, Dr. Jennifer Wilcox, Dr.

 

 

xvii  

Jason Stephens, Kaitlin Haas, Erin Cullen, and Natalie Lamberton. My partner in crime,

Dr. Joy Gallagher, has always been there to listen and provide her perspective on my

research and my life. From our lunchtime study sessions in our first year to thesis review

sessions in our last few months, she has been an excellent confidant and provided endless

entertainment. My officemate, Dr. Jennifer Wilcox, has been my support in these last few

months. Together we have managed our stress, and I cannot think of a person I would

rather share my space or my snacks with.

I am fortunate the have the unwavering support of a wonderful family that has

been a source of incredible strength throughout my life. My father, Joseph Zaccardi, has

always given me his support and encouragement, and made me feel like I could tackle

seemingly insurmountable tasks. My mother, Joan Zaccardi, has been a sounding board

for my frustrations, and has provided great advice even when I did not think I needed it.

My siblings, Diane Baldwin and Joseph Zaccardi III, have been role models for me

throughout my life, and are excellent examples in attaining your goals. Lastly, I would

like to thank the greatest part of my life, my dog, Grace. She has learned more science

from holding my hand and listening as I practice all of my presentations that she probably

deserves her own dog doctorate. Taking care of her also forced me to take a break and get

some exercise on the busiest and most stressful days, allowing me to refocus and rest.

“This seems to be the law of progress in everything we do: it moves along a spiral

rather than a perpendicular; we seem to be actually going out of the way, and yet it

turns out that we were really moving upward all the time.”

Frances E. Willard

 

 

1  

Chapter 1

Introduction to Indole-3-glycerol Phosphate Synthase

1.1 Progress Towards Engineering Enzymes with New Functions

Biocatalysts offer a unique solution to many common industrial synthesis

problems due to their high efficiencies and high degree of selectivity. They also allow for

the use of mild reaction conditions compared to typical industrial processes.1, 2 Because

of these properties, enzymes are being increasingly implemented in a variety of industries

including bioremediation, textiles, biofuel production, pharmaceuticals, and agriculture.3

For example, an optimized lipase is used to synthesize enantiomer specific precursors for

the production of diltiazem, a blood pressure medication.4 Much of the current industrial

application of enzymes has focused on improving stability, or changing specificity for

reactions already performed in natural enzyme systems. This focus is the result of a

desire to utilize biological systems amongst those industrial processes that have already

been optimized under conditions not typically suited for enzymes, such as high

temperatures, extreme pHs, high concentrations, and non-aqueous solvents.5 However,

there is also value in the production of “made to order” enzymes that can catalyze

reactions not catalyzed in nature; for many industrially important syntheses, a naturally

occurring enzyme capable of catalyzing the reaction does not exist.4, 6 Engineering

enzymes capable of catalyzing non-natural reactions at rates comparable to enzymes

found in nature challenges the fundamental understanding of enzyme function. In order to

recreate the mechanisms evolved in biological systems, we must completely understand

 

 

2  

how sequence, structure, conformational dynamics, and function are intertwined, and

work together in catalyzing the reaction.

Over the last several years, a large number of studies have focused on the

synthesis of novel enzymes.7-17 The explosion in research on this subject largely began

with two renowned studies from the Baker laboratory.14, 15 In these studies, two non-

natural reactions were engineered onto natural enzyme scaffolds. First, plausible

chemical mechanisms along with appropriate transition states were established and then

computational methods were used to create an active site with the appropriate

architecture, using several different enzyme structures as a starting scaffold including the

(β/α)8-barrel and the jelly roll. In the first study, a retro-aldol enzyme capable of breaking

a carbon-carbon bond of a non-natural substrate was designed.14 The second study

engineered an enzyme for the Kemp elimination reaction, which requires the direct

removal of a hydrogen from a carbon, a process that is not possible through normal

synthetic routes due to its high activation energy barriers.15 While both studies were able

to engineer enzymes with an enhancement of the desired activity on the order of 106 for

the Kemp elimination and 104 for the retro-aldol reaction, these rate enhancements are

still very low compared to the rates of natural enzymes.14, 17 Reoptimization of both

systems has also been performed.17, 18 For the Kemp elimination, considerable increase in

catalytic rates was possible through directed evolution of the previously designed

catalysts. However, directed evolution studies are very time intensive, require screening

of a large number of inactive variants, and were still unable to match the rate

enhancement of biological systems.16

 

 

3  

These studies are largely based on modeling the active site architecture, and

despite the high similarity of the crystal structures on the non-natural enzymes compared

to their models, reaction rate enhancements (105 to 106) are not able to match natural

enzymes (1023).19 This result may be because the algorithms used to design these systems

only integrate catalytically relevant residues, and disregard residues that may be

important for other processes. For example, molecular dynamics (MD) simulations show

that the enzymes designed for the Kemp elimination have dynamic fluctuations that

prevent proper active site configuration, and thus the enzymes show lower than expected

reaction rates.20 This finding highlights the need for enzyme engineering studies to also

take into account protein motions, and as such, many researchers have introduced

dynamics into their design algorithms. This advancement has allowed for some

improvements in rate enhancement, although a more complete understanding of how

sequence changes affect dynamic processes will be beneficial for the further development

of these techniques.20-22

Although enzymes are regularly depicted as rigid structures as they are found in

crystals, in solution enzymes are quite flexible, and undergo conformational fluctuations

that are thought to be important for regulating substrate binding, catalysis, folding, and

other processes. The role of flexibility in enzyme function has been widely studied, and

its importance in catalysis has been implicated in many different enzymes.23-29 However,

even with the growing knowledge of enzyme dynamics, reaction rates of engineered

enzymes were only enhanced by about five orders of magnitude over the rate of the

uncatalyzed reaction, whereas reaction rates of natural enzymes can be enhanced up to

twenty-three orders of magnitude.

 

 

4  

Engineering new enzymes that are capable of performing non-natural reactions

will provide significant improvements to the technology in many different industries.

Industrial synthetic processes are typically harsher than those found in nature, and often

utilize high temperatures, extreme pH conditions, and non-aqueous solvents. The ability

to introduce enzymes that can withstand these extremes, or decrease the need for such

conditions, will drastically improve the efficiency of industrial processes.5 However, in

order to produce viable enzymes for these applications, a fundamental understanding of

the molecular determinants not just for catalysis, but also for structure, stability, and

dynamics is required. The process of changing the active site architecture from one

enzyme to another is intricate, and requires a very detailed understanding of all the

factors that can affect activity in the new enzyme including protein folding, flexibility,

stability, structure, and chemical interactions.

In order to engineer enzymes capable of catalyzing non-natural reactions at rates

closer to those found in nature, a more comprehensive approach is required that takes into

account not only the residues required for the new activity, but also the starting enzyme

architecture, and the role of both active site residues and residues more distant from the

active site in the scaffold enzyme. This undertaking requires an in depth knowledge for

the role of all residues in the protein, especially considering that semi-conserved or non-

conserved residues, in addition to those that are conserved, contribute to active site

architecture, affect the ability of an enzyme to catalyze the desired reaction, and are

involved in other processes like protein folding and stability.3

 

 

5  

1.2 The Conserved (β/α)8-Barrel Protein Fold as an Enzyme Engineering Scaffold

Enzymes containing the (β/α)8-barrel (or TIM-barrel, named after triose

phosphate isomerase) fold have been widely used as enzyme engineering scaffolds due to

the fold’s high stability, diverse catalytic ability, and conserved structure. In fact, the

(β/α)8-barrel shows higher success in these studies (including the Kemp elimination and

retro aldolase reactions previously described) than other folds.14,15 The (β/α)8-barrel is the

most common enzyme fold in nature and is found in many different enzyme

superfamilies that are capable of catalyzing a diverse range of reactions.30-32 It generally

consists of eight units of alternating β-strands and α-helices that are connected by loops.

The β-strands form a barrel in the center and are surrounded by the α-helices. This

dualism of conserved structure with diverse function of (β/α)8-barrel enzymes provides an

excellent model for studying the relationship between enzyme sequence, structure,

function, and dynamics. Catalytically important residues are typically found on the C-

terminal ends of the β-sheets, and on the loops connecting the β-strands to the α-helices

(βα loops).33 This structure provides several different “take off” positions in enzyme

engineering studies that can be used for catalytic residues and transition state

stabilization, as all positions pointing towards the inside of the barrel can be used.15

The role of dynamics in enzymes containing the (β/α)8-barrel fold has been well

documented.34-36 In the iconic (β/α)8-barrel, triosephosphate isomerase (TIM), several

studies have examined loop dynamics that contribute to the catalysis by the enzyme.37-40

The β6α6 loop in TIM undergoes conformational exchange to allow for substrate binding

and product release. Similar behavior is seen for other (β/α)8-barrel enzymes including

alkanesulfaonate monooxygenase,41 imidazole glycerol phosphate synthase,42 and D-

 

 

6  

ribulose 5-phosphate 3-epimerase.43 A better understanding of how specific amino acids

change the conformational dynamics of these enzymes would improve the ability to fine

tune the (β/α)8-barrel scaffold for new functions.

1.3 (β/α)8-Barrel Enzymes in Tryptophan Biosynthesis

In tryptophan biosynthesis, there are three consecutive enzymes that contain this

(β/α)8-barrel fold: N-(5’-phosphoribosyl)-anthranilate isomerase (PRAI), indole-3-

glycerol phosphate synthase (IGPS), and the alpha subunit of tryptophan synthase (αTS)

(Figure 1.1), which catalyze the fourth, fifth, and sixth committed steps of tryptophan

production.44, 45 PRAI catalyzes an Amadori rearrangement of phosphoribosyl

anthranilate (PRA) to form 1-(o-carboxyphenylamino)-1-deoxyribulose 5-phosphate

(CdRP). IGPS then performs a ring closure to form indole-3-glycerol phosphate (IGP).

Lastly, αTS removes the glyceraldehyde 3-phosphate from IGP to form the indole ring

that goes forward in the pathway and combines with L-serine to form tryptophan. It is

interesting from an evolutionary standpoint that these three enzymes catalyze different

types of reactions but all evolved with the same fold, leading to several possible

hypotheses for the evolution of the tryptophan biosynthetic pathway. First, all three

enzymes may have evolved divergently from a common ancestral enzyme that was

capable of catalyzing all three reactions with poor efficiency.30, 31, 46 In the divergent

mechanism, the enzymes may have evolved through gene duplication, where the

organism made multiple copies of a gene that could then evolve several different

functions.47, 48

 

 

7  

Figure 1.1: The three consecutive (β/α)8-barrel enzymes in tryptophan biosythnesis catalyze the fourth, fifth, and sixth steps of the pathway. PRAI (PDB 1Pll) catalyzes the amadori rearrangement of PRA to form CdRP. CdRP is then converted to IGP by IGPS (PDB 1Pll). Then, αTS (PDB 1V7Y) catalyzes the cleavage of IGP to form glyceraldehyde-3-phosphate and indole.

 

 

8  

Second, akin to proposals by Horowitz regarding the evolution of metabolism, the

pathway may have evolved backwards, with αTS evolving first, followed by IGPS, and

lastly PRAI.49 Several studies have explored the relationship between related (β/α)8-barrel

enzymes.47, 50-52 One such directed evolution study by Evran et al. was able to establish

PRAI activity using the α-TS scaffold.51 This study provided evidence of evolution of the

(β/α)8-barrel enzymes in tryptophan synthesis by divergence/gene duplication.52 Despite

the low sequence identity of these three enzymes (PRAI and IGPS are only about 22%

sequence identical, and αTS shows even lower sequence identity with both PRAI and

IGPS), they contain very conserved structural elements, including the phosphate binding

site located in the β7α7 and β8α8 loops. However, other loops have been shown to be

important for specific activity, as in one study in which loop swapping between PRAI

and α-TS changed the catalytic ability of the enzyme.34, 53

1.4 The Tryptophan Biosynthetic Enzyme Indole-3-glycerol Phosphate Synthase

In general the (β/α)8-barrel fold has shown greater success as an engineering

scaffold than other folds, and in several studies the largest number of active variants, as

well as those with the highest rate enhancement, were produced using IGPS from

Sulfolobus sulfataricus (ssIGPS) as a scaffold.14, 15 In fact, ssIGPS was used as a starting

scaffold in the studies from the Baker laboratory, and show the highest rate enhancement

compared to all other enzyme scaffolds examined (including TIM).14, 15 In addition to the

promise ssIGPS shows in novel enzyme design, its naturally catalyzed reaction is also

applicable for the industrial synthesis of indole and its derivatives. ssIGPS is a

thermophilic enzyme; therefore, its increased stability is desireable for industrial

 

 

9  

processes, and the indole ring is a widely used structure in pharmaceuticals, agriculture,

and other industries.54

IGPS is also valuable in several other fields including the development of new

antimicrobial agents and the understanding of thermophilic enzymes.55 The tryptophan

biosynthetic pathway is not found in humans, but is found in pathogenic bacteria.

Therefore, the enzymes in the pathway are potential targets for new antimicrobial

compounds, particularly those pathogens that show a high occurance of multidrug

resistance such as Mycobacterium tuberculosis. Due to the widespread multidrug

resistance of the bacterium to currently available treatments, as well as the prevalence of

tuberculosis (TB) in impoverished areas of the world, the disease can be difficult to treat

and the currently available treatments are expensive, complex, and can have harsh side

effects.55 These characteristics lead to poor patient compliance and a push to identify new

targets for effective TB treatment.56

Studies by Smith et al. demonstrate that tryptophan auxotrophs of M. tuberculosis

were avirulent in mice, indicating that the bacterium may be unable to uptake these

amino acids in vivo.57 The gene that encodes for the IGPS enzyme (TrpC) was also

shown to be essential for growth of M. tuberculosis in vitro,58 which further demonstrates

the potential utility in targeting IGPS as a treatment for TB and other microbial diseases.

While there are several known inhibitors for IGPS,55, 59, 60 none are currently available for

treatment of tuberculosis or other bacterial infections. Studies towards the development

of new antimicrobials that target IGPS will benefit from a more in depth examination of

its mechanism and active site.

 

 

10  

Thermophilic organisms have evolved robust mechanisms to overcome the

deleterious effects high temperature typically has on biomolecules, such as denaturation,

allowing life to exist under extreme conditions, including extreme temperatures. The

thermophilic archaeon, S. sulfolobus, is found at high temperatures (> 80 °C) and has

developed mechanisms in order to exist at extreme temperatures. For enzymes, this

includes an increase in stabilizing interactions such as hydrogen bonds and electrostatic

interactions, as well as higher packing efficiencies and increased burial of the

hydrophobic surfaces.61-64 Harnessing the stabilizing properties of thermophilic enzymes

without disrupting catalytic activity will provide major technological advancements for

the development of enzymes for biotechnology industries.

A fuller understanding of IGPS would also be useful for understanding

temperature adaptation, as IGPS has been widely studied in both thermophilic and

mesophilic organisms. Crystal structures and biochemical techniques have identified

structural differences between IGPS from S. sulfataricus and E. coli (ecIGPS),65, 66 as

well as Thermotoga maritima (tmIGPS),67 Thermatoga thermophilus,68 and

Thermococcus kadakarensis.69 Despite their high structural similarity, ssIGPS and

ecIGPS are only about 30% sequence identical, and ssIGPS contains additional

noncovalent interactions that prevent its denaturation at increased temperatures. ssIGPS

also shows decreased activity at lower temperatures compared to its mesophilic

counterpart in ecIGPS.64, 65, 70 Some studies suggest that the temperature dependent

activity differences between thermophilic ssIGPS and mesophilic ecIGPS is the result of

a decrease in flexibility of the protein caused by the stabilizers (e.g. salt bridges) that are

needed to adapt to the increase in temperature.71, 72 Indeed, Merz et al. semiquantitatively

 

 

11  

examined the flexibility of ssIGPS at various temperatures through a limited proteolysis

study, and the results showed the less flexible the IGPS enzyme at lower temperatures,

the lower its rate of thermal inactivation, and the lower its activity at lower

temperatures.64

Despite the differences in activity for ssIGPS at lower versus higher temperatures,

little research has been performed on ssIGPS near its biologically relevant temperatures.

Studies on ssIGPS at lower temperatures suggest that the rate-determining step of the

overall reaction is product release.64, 73 However, there may be differences between the

kinetic or chemical reaction mechanism at lower versus higher temperatures as well as

between thermophilic and mesophilic homologs. A better understanding of catalysis for

both thermophiles and mesophiles over a range of temperatures will aid in the application

of the robust and stable enzymes in an industrial setting.

1.3 Previous Knowledge on the Mechanism of IGPS

The mechanism for the conversion of CdRP to form IGP by IGPS (Figure 1.2)

was originally proposed nearly forty years ago by Parry et al. The reaction is proposed to

occur in three steps (condensation, decarboxylation, and dehydration) with two distinct

intermediates (I1 and I2).74 This mechanism, particularly the formation of I1, was

motivated by the observation that the reaction does not occur for a substrate analog

lacking the carboxyl, which is evidence that the carboxyl is required for pyrrole ring

formation.75

 

 

12  

Figure 1.2: Indole-3-glyerol phosphate synthase. (a) ssIGPS (PDB: 1IGP) is a (β/α)8-barrel enzyme that contains an additional 45 residue N-terminal extension compared to the standard (β/α)8-barrel fold. (b) IGPS catalyzes the conversion of CdRP to form IGP. The proposed mechanism contains three steps and two intermediates and utilizes a general acid and base (proposed as Lys110 and Glu159).74,77

 

 

13  

Multiple sequence alignment of IGPS from various species shows several

conserved residues (Figure 1.3). Further insight into the possible role of these residues

and the mechanism of IGPS came from studies of amino acid substitutions in IGPS from

ecIGPS including Glu53, Lys55, Lys114, Glu163, Asn184, and Arg186 (corresponding

to Glu51, Lys53, Lys110, Glu159, Asn180, Arg182 in ssIGPS).76 The proposed general

acid, Lys114, is essential, and neither Arg nor His amino acid substitutions in this

position yielded active enzyme. Similar results were found for the Glu163Asp variant,

which lead to the assignment of Lys114 and Glu163 as the general acid-base pair in

ecIGPS. Amino acid substitutions at Lys55 and Glu53 also yielded interesting results,

with a 40-fold decrease in kcat for the Glu53Cys variant. This finding indicates that Glu53

is important for catalysis, and the authors suggest it may help with the proper positioning

of Lys114. The Lys55Ser variant shows a twenty-fold decrease in kcat and a 1800-fold

increase in KM, indicating a role for this residue in both chemistry and ligand binding.

While these experiments identified those residues that are catalytically required, they

were unable to discern the step of the reaction in which the residues were involved.

Additional insight about IGPS was gained from crystal structures of IGPS bound

with substrate, substrate analog, and product, which suggested roles for several amino

acids in IGPS catalysis (Figure 1.4, Table 1.1), including the proposed general acid and

base, Lys110 and Glu159, respectively (numbering according to ssIGPS).77 Arg182 and

Phe89 were predicted to be involved in substrate binding with Phe89 interacting with the

aromatic moiety and Arg182 interacting with the phosphate group. The roles for Glu51

and Lys53 were more ambiguous. The authors asserted that Lys53 helps bind the

substrate, but that Glu51 and Lys53 can also form a salt bridge triad with Lys110 that

 

 

14  

Figure 1.3: Multiple sequence alignments between IGPS from S. sulfataricus (ssIGPS), Thermatoga maritima (tmIGPS), E. coli (ecIGPS), and Mycobacterium tuberculosis (mtIGPS). Secondary structure is denoted by boxes above the sequence with α-helices in green and β-sheets in blue. Conserved residues are bold and in blue. Catalytically relevant residues are denoted with a star. Sequence alignment was performed with Clustal Omega provided by The European Bioinformatics Institute at The European Molecular Biology Laboratory.

 

 

15  

Figure 1.4: Active site of IGPS with reduced CdRP bound. Conserved and catalytically relevant residues are shown. Lys110 is the proposed general acid in the condensation and dehydration steps. Glu159 and Glu210 have both been proposed to act as general base. Phe89 and Arg182 are proposed to aid in substrate binding. Lys53 is also involved in substrate binding and may have additional roles in the chemical steps. The role of Glu51 has not been extensively studied.76,77

 

 

16  

Table 1.1: Conserved active site residues in ssIGPS that are of interest to these studies are shown along with their proposed roles in enzyme activity.76,77,78

Conserved Residue Location in ssIGPS Proposed Role Glu51 β1 Strand Interacts with Lys110 and Lys53 Lys53 β1α1 Loop Substrate binding Phe89 β2α2 Loop Substrate binding Lys110 β3 Strand General acid Glu159 β5 Strand General base

(predicted by crystal structures) Arg182 β6α6 Loop Substrate binding Glu210 β7 Strand General base

(predicted by MD simulations)

 

 

17  

may aid the activity of the general acid. Additionally, the crystal structure showed the

anthranilate group bound into a different hydrophobic pocket in IGPS complexed with

substrate than when complexed with product (Figure 1.5), introducing the idea that the

substrate may undergo conformational rearrangement in the active site during catalysis.

Lys53 is conserved and is located on the highly dynamic β1α1 loop. Crystal

structures predict that it is largely involved in substrate binding, as it can hydrogen bond

in multiple positions of CdRP including the C1 carboxyl and the C3’ hydroxyl

(numbering for CdRP, Figure 1.6). MD studies suggest Lys53 is important for structural

rearrangements that facilitate catalysis, and have suggested that the interaction of Lys53

with the substrate along with the flexibility of the β1α1 loop, may be involved in bridging

the gap between the C1 and C2’ allowing for the formation of the pyrrole ring.70, 78 The

authors asserted that enzyme flexibility is required for the formation of a near attack

conformer (NAC). NACs are groundstate substrate conformations that can convert most

efficiently into the transition state.78 For CdRP in the IGPS catalyzes reaction, the NAC is

defined by the reacting moeties, C1-C2’, within van der Waals contact distance (≤ 3.5 Å)

and at an approaching angle of 120° ± 20°.70,78 Similarly, work performed by Goodey and

Sterner indicates that the flexibility of this loop is coupled to enzyme activity.73

Other studies further highlight the potential role for dynamics and flexibility in

IGPS. Shen et al. examined correlated and coevolving residues in IGPS through a

combined study using statistical coupling analysis (SCA) and MD simulations.79 SCA

identifies residues that are not necessarily conserved, but covary between the enzymes

from different species. This method identified amino acid pairs whose interaction may be

important for protein folding, stability, or catalysis.80, 81 When combined with MD

 

 

18  

Figure 1.5: IGPS in complex with rCdRP (PDB:1LBF) (yellow) and IGP (1A53) (blue) showing residues that interact with the aromatic, anthranilate moiety. When CdRP binds in the active site, Trp8, Pro57, Phe89, Arg182, and Leu184 interact with the anthranilate. Conversely, when IGP binds, Phe89, Lys110, Phe112, Ile133, and Arg182 interact.

 

 

19  

Figure 1.6: Numbering for CdRP. Lys53 interacts with the C1 carboxyl and C3’ hydroxyl groups and is thought to aid in ring closure between C1 and C2’.

 

 

20  

simulations, this study identified amino acid pairs in IGPS that showed the potential

importance of residues that are not necessarily conserved, but whose coordinated motion

is potentially important in regulating a variety of enzyme processes including

conformational exchange, enzyme folding, structural stability, or chemistry itself.82 The

authors assert that these amino acid pairs, which are all in van der Waals contact with one

another, form an amino acid network throughout the enzyme that aids in enzyme activity.

Most notable to this work are active site residues Arg54, Glu85, and Asn90. Their

location on dynamic active site loops, and their proximity to conserved and catalytic

residues may be indicative of a more direct role, potentially important for coordinating

functional motion in IGPS catalysis.

1.6 Conclusions

Considering the potential application for the IGPS enzyme across many different

industries, it is essential that the mechanism of IGPS be very well understood. For many

of these applications, a detailed understanding of the role of active site residues in the

catalytic mechanism is required, especially considering that a single mutation or small

structural change can cause large differences in enzyme function, folding, and/or

stability.36 In order to apply IGPS as an industrial enzyme, use it as a target for

antibacterial agents, or as a scaffold for enzyme engineering, we must understand all of

the active site residues, not just those that are conserved, but also those residues that may

contribute to enzyme architecture, dynamics, and other processes even if they are distal

from the active site. Even residues that are not conserved can play a large role in the

catalytic mechanism, or participate in other processes such as protein folding, stability, or

 

 

21  

dynamics. Failure to consider how an enzyme undergoes catalysis will lead to less

efficient application of IGPS for enzyme engineering and as a antimicrobial target.

Despite the extensive research on IGPS, there are still many questions regarding

the kinetic and chemical mechanism as well as the specific role of several conserved

residues in catalysis. First, previous research only examined the kinetic mechanism of

ssIGPS at lower temperatures (25 °C). In Chapter 2, the kinetic differences between

thermophilic ssIGPS and mesophilic ecIGPS are examined. The results show that the

kinetic mechanism of ssIGPS is temperature dependent, with product release being rate-

determining at lower temperatures (25 °C) and the ring closure being rate-determining at

higher, biologically relevant temperatures (75 °C). Additionally, ssIGPS and ecIGPS

display different rate-determining steps at their adaptive temperatures.

Another issue with the mechanism of IGPS involves the general acid, Lys110,

which is suggested to donate a proton in both the condensation and dehydration steps,

although there has been no plausible mechanism suggested for its reprotonation. Several

studies are also at odds in the assignment of the general base, with crystallography

suggesting Glu159 performs this task and MD simulations suggesting Glu210. There are

also several other conserved residues in the active site whose roles are undefined. In

Chapter 3, the role of conserved, charged active site residues are examined. The results

show that Lys53 and Glu51 are the general acid and base in the dehydration step. This

finding is at odds with the previously published mechanism for IGPS that suggested the

general acid base pair was Lys110/Glu159. This new assignment also led to the proposal

that the substrate undergoes a reorientation in the active site after the first chemical step

 

 

22  

in order to be properly aligned for catalysis. This study also suggests that the substrate

must undergo a reorientation in the active site.

Lastly, several studies have implicated a role for active site loops in catalysis

including computational research suggesting a functional role for coevolving residues in

catalysis was unspecific.78 A better understanding of the role for coevolving residues in

the IGPS is desirable for future applications. Therefore, in Chapter 4, the role of

coevolving, active site residues Arg54, Glu85, and Asn90 were examined. These residues

are not conserved, but our results suggest that they are still involved in proper function of

ssIGPS. Specifically, the interaction between Arg54 and Asn90 is involved in the proper

function of the general acid and base, Lys53 and Glu51, during dehydration. The research

presented in this dissertation has more completely examined the kinetic and chemical

mechanism for IGPS and has greatly improved the understanding of catalysis by this

enzyme.

1.6 References

1.   Nestl,   B.   M.;   Nebel,   B.   A.;   Hauer,   B.,   Recent   progress   in   industrial   biocatalysis.  Current  Opinion  in  Chemical  Biology  15  (2),  187-­‐193.    2.   Wang,   M.;   Si,   T.;   Zhao,   H.,   Biocatalyst   development   by   directed   evolution.  Bioresource  Technology  115,  117-­‐125.    3.   Kiss,   G.;   Roethlisberger,   D.;   Baker,   D.;   Houk,   K.   N.,   Evaluation   and   ranking   of  enzyme  designs.  Protein  Science  2010,  19  (9),  1760-­‐1773.    4.  Bornscheuer,  U.  T.;  Huisman,  G.  W.;  Kazlauskas,  R.  J.;  Lutz,  S.;  Moore,  J.  C.;  Robins,  K.,  Engineering  the  third  wave  of  biocatalysis.  Nature  2012,  485  (7397),  185-­‐194.    5.  Woodley,   J.  M.,  Protein  engineering  of  enzymes  for  process  applications.  Current  Opinion  in  Chemical  Biology  2013,  17  (2),  310-­‐316.    

 

 

23  

6.  Campeotto,  I.;  Acevedo-­‐Rocha,  C.  G.,  Hijacking  nature-­‐new  approaches  to  unravel  enzyme  mechanisms   and   engineer   improved   biocatalysts.  Embo  Reports  2013,  14  (4),  299-­‐301.      7.  Linder,  M.;  Johansson,  A.  J.;  Olsson,  T.  S.  G.;  Liebeschuetz,  J.;  Brinck,  T.,  Designing  a  New   Diels-­‐Alderase:   A   Combinatorial,   Semirational   Approach   Including   Dynamic  Optimization.   Journal   of   Chemical   Information   and   Modeling   2011,   51   (8),   1906-­‐1917.    8.   Alvizo,   O.;   Mittal,   S.;   Mayo,   S.   L.;   Schiffer,   C.   A.,   Structural,   kinetic,   and  thermodynamic   studies   of   specificity   designed   HIV-­‐1   protease.   Protein   Science  2012,  21  (7),  1029-­‐1041.    9.  Chen,  M.  M.  Y.;  Snow,  C.  D.;  Vizcarra,  C.  L.;  Mayo,  S.  L.;  Arnold,  F.  H.,  Comparison  of  random  mutagenesis  and  semi-­‐rational  designed  libraries  for  improved  cytochrome  P450  BM3-­‐catalyzed  hydroxylation   of   small   alkanes.  Protein   Engineering  Design  &  Selection  2012,  25  (4),  171-­‐178.    10.  Privett,  H.  K.;  Kiss,  G.;  Lee,  T.  M.;  Blomberg,  R.;  Chica,  R.  A.;  Thomas,  L.  M.;  Hilvert,  D.;   Houk,   K.   N.;   Mayo,   S.   L.,   Iterative   approach   to   computational   enzyme   design.  Proceedings   of   the   National   Academy   of   Sciences   of   the   United   States   of   America  2012,  109  (10),  3790-­‐3795.    11.  Smith,  C.  A.;  Shi,  C.  A.;  Chroust,  M.  K.;  Bliska,  T.  E.;  Kelly,  M.  J.  S.;  Jacobson,  M.  P.;  Kortemme,   T.,   Design   of   a   Phosphorylatable   PDZ   Domain   with   Peptide-­‐Specific  Affinity  Changes.  Structure  2013,  21  (1),  54-­‐64.    12.  Karanicolas,  J.;  Com,  J.  E.;  Chen,  I.;  Joachimiak,  L.  A.;  Dym,  O.;  Peck,  S.  H.;  Albeck,  S.;  Unger,  T.;  Hu,  W.;  Liu,  G.;  Delbecq,  S.;  Montelione,  G.  T.;  Spiegel,  C.  P.;  Liu,  D.  R.;  Baker,  D.,   A  De  Novo  Protein  Binding   Pair   By   Computational  Design   and  Directed  Evolution.  Molecular  Cell  2011,  42  (2),  250-­‐260.    13.   Linder,   M.;   Johansson,   A.   J.;   Olsson,   T.   S.   G.;   Liebeschuetz,   J.;   Brinck,   T.,  Computational   design   of   a   Diels-­‐Alderase   from   a   thermophilic   esterase:   the  importance  of  dynamics.  Journal  of  Computer-­Aided  Molecular  Design  2012,  26  (9),  1079-­‐1095.    14.  Jiang,  L.;  Althoff,  E.  A.;  Clemente,  F.  R.;  Doyle,  L.;  Rothlisberger,  D.;  Zanghellini,  A.;  Gallaher,  J.  L.;  Betker,  J.  L.;  Tanaka,  F.;  Barbas,  C.  F.;  Hilvert,  D.;  Houk,  K.  N.;  Stoddard,  B.  L.;  Baker,  D.,  De  novo  computational  design  of  retro-­‐aldol  enzymes.  Science  2008,  319  (5868),  1387-­‐1391.    15.  Rothlisberger,  D.;  Khersonsky,  O.;  Wollacott,  A.  M.;  Jiang,  L.;  DeChancie,  J.;  Betker,  J.;  Gallaher,  J.  L.;  Althoff,  E.  A.;  Zanghellini,  A.;  Dym,  O.;  Albeck,  S.;  Houk,  K.  N.;  Tawfik,  

 

 

24  

D.  S.;  Baker,  D.,  Kemp  elimination  catalysts  by  computational  enzyme  design.  Nature  2008,  453  (7192),  190-­‐U4.      16.   Khersonsky,   O.;   Rothlisberger,   D.;   Dym,   O.;   Albeck,   S.;   Jackson,   C.   J.;   Baker,   D.;  Tawfik,   D.   S.,   Evolutionary   Optimization   of   Computationally   Designed   Enzymes:  Kemp   Eliminases   of   the   KE07   Series.   Journal   of   Molecular   Biology  2010,   396   (4),  1025-­‐1042.    17.  Khersonsky,  O.;  Roethlisberger,  D.;  Wollacott,  A.  M.;  Murphy,  P.;  Dym,  O.;  Albeck,  S.;  Kiss,  G.;  Houk,  K.  N.;  Baker,  D.;  Tawfik,  D.  S.,  Optimization  of  the  In-­‐Silico-­‐Designed  Kemp  Eliminase  KE70  by  Computational  Design  and  Directed  Evolution.   Journal  of  Molecular  Biology  2011,  407  (3),  391-­‐412.    18.  Althoff,  E.  A.;  Wang,  L.;  Jiang,  L.;  Giger,  L.;  Lassila,  J.  K.;  Wang,  Z.;  Smith,  M.;  Hari,  S.;  Kast,  P.;  Herschlag,  D.;  Hilvert,  D.;  Baker,  D.,  Robust  design  and  optimization  of  retroaldol  enzymes.  Protein  Science  21  (5),  717-­‐726.    19.  Walsh,  C.,  Enabling  the  chemistry  of  life.  Nature  2001,  409,  226-­‐231.    20.   Ruscio,   J.   Z.;   Kohn,   J.   E.;   Ball,   K.   A.;   Head-­‐Gordon,   T.,   The   Influence   of   Protein  Dynamics  on  the  Success  of  Computational  Enzyme  Design.  Journal  of  the  American  Chemical  Society  2009,  131  (39),  14111-­‐14115.    21.  Correia,  B.  E.;  Ban,  Y.-­‐E.  A.;  Friend,  D.   J.;  Ellingson,  K.;  Xu,  H.;  Boni,  E.;  Bradley-­‐Hewitt,   T.;   Bruhn-­‐Johannsen,   J.   F.;   Stamatatos,   L.;   Strong,   R.   K.;   Schiefl,   W.   R.,  Computational   Protein   Design   Using   Flexible   Backbone   Remodeling   and  Resurfacing:   Case   Studies   in   Structure-­‐Based  Antigen  Design.   Journal   of  Molecular  Biology  2011,  405  (1),  284-­‐297.    22.   Lassila,   J.   K.,   Conformational   diversity   and   computational   enzyme   design.  Current  Opinion  in  Chemical  Biology  2010,  14  (5),  676-­‐682.    23.   Ramanathan,   A.;   Agarwal,   P.   K.,   Evolutionarily   Conserved   Linkage   between  Enzyme  Fold,  Flexibility,  and  Catalysis.  Plos  Biology  2011,  9  (11).    24.  Boehr,  D.  D.;  Dyson,  H.  J.;  Wright,  P.  E.,  An  NMR  perspective  on  enzyme  dynamics.  Chemical  Reviews  2006,  106  (8),  3055-­‐3079.    25.   Boehr,   D.   D.;   Nussinov,   R.;   Wright,   P.   E.,   The   role   of   dynamic   conformational  ensembles   in   biomolecular   recognition   (vol   5,   pg   789,   2009).   Nature   Chemical  Biology  2009,  5  (12),  954-­‐954.    26.   Boehr,   D.   D.;   McElheny,   D.;   Dyson,   H.   J.;   Wright,   P.   E.,   The   dynamic   energy  landscape   of   dihydrofolate   reductase   catalysis.   Science   2006,   313   (5793),   1638-­‐1642.  

 

 

25  

27.   Hammes,   G.   G.,   Multiple   conformational   changes   in   enzyme   catalysis.  Biochemistry  2002,  41  (26),  8221-­‐8228.  28.   Kempner,   E.   S.,   MOVABLE   LOBES   AND   FLEXIBLE   LOOPS   IN   PROTEINS   -­‐  STRUCTURAL   DEFORMATIONS   THAT   CONTROL   BIOCHEMICAL-­‐ACTIVITY.   Febs  Letters  1993,  326  (1-­‐3),  4-­‐10.    29.   Skliros,   A.;   Zimmermann,   M.   T.;   Chakraborty,   D.;   Saraswathi,   S.;   Katebi,   A.   R.;  Leelananda,  S.  P.;  Kloczkowski,  A.;   Jernigan,  R.  L.,  The   importance  of   slow  motions  for  protein  functional  loops.  Physical  Biology  2012,  9  (1).    30.   Gerlt,   J.   A.;   Babbitt,   P.   C.,   Divergent   evolution   of   enzymatic   function:  Mechanistically   diverse   superfamilies   and   functionally   distinct   suprafamilies.  Annual  Review  of  Biochemistry  2001,  70,  209-­‐246.    31.  Copley,  R.  R.;  Bork,  P.,  Homology  among  (beta  alpha)(8)  barrels:  Implications  for  the   evolution   of  metabolic   pathways.   Journal   of   Molecular   Biology  2000,   303   (4),  627-­‐640.    32.  Vega,  M.  C.;  Lorentzen,  E.;  Linden,  A.;  Wilmanns,  M.,  Evolutionary  markers  in  the  (beta/alpha)8-­‐barrel  fold.  Curr  Opin  Chem  Biol  2003,  7  (6),  694-­‐701.    33.   Reisinger,   B.;   Bocola,  M.;   List,   F.;   Claren,   J.;   Rajendran,   C.;   Sterner,   R.,   A   sugar  isomerization   reaction   established   on   various   (beta   alpha)(8)-­‐barrel   scaffolds   is  based  on  substrate-­‐assisted  catalysis.  Protein  Engineering  Design  &  Selection  2012,  25  (11),  751-­‐760.    34.  Ochoa-­‐Leyva,  A.;  Barona-­‐Gomez,  F.;  Saab-­‐Rincon,  G.;  Verdel-­‐Aranda,  K.;  Sanchez,  F.;   Soberon,   X.,   Exploring   the   Structure-­‐Function   Loop   Adaptability   of   a  (beta/alpha)(8)-­‐Barrel   Enzyme   through   Loop   Swapping   and   Hinge   Variability.  Journal  of  Molecular  Biology  2011,  411  (1),  143-­‐157.    35.  Malabanan,  M.  M.;  Amyes,  T.  L.;  Richard,  J.  P.,  A  role  for  flexible  loops  in  enzyme  catalysis.  Current  Opinion  in  Structural  Biology  2010,  20  (6),  702-­‐710.    36.  Glembo,  T.   J.;   Farrell,  D.  W.;  Gerek,   Z.  N.;  Thorpe,  M.  F.;  Ozkan,   S.  B.,   Collective  Dynamics   Differentiates   Functional   Divergence   in   Protein   Evolution.   Plos  Computational  Biology  2012,  8  (3).    37.   Rozovsky,   S.;  McDermott,   A.   E.,   The   time   scale   of   the   catalytic   loop  motion   in  triosephosphate  isomerase.  Journal  of  Molecular  Biology  2001,  310  (1),  259-­‐270.    38.   Massi,   F.;  Wang,   C.;   Palmer,   A.   G.,   III,   Solution   NMR   and   computer   simulation  studies  of  active  site  loop  motion  in  triosephosphate  isomerase.  Biochemistry  2006,  45  (36),  10787-­‐10794.    

 

 

26  

39.   Rozovsky,   S.;   Jogl,   G.;   Tong,   L.;   McDermott,   A.   E.,   Solution-­‐state   NMR  investigations  of  triosephosphate  isomerase  active  site  loop  motion:  Ligand  release  in  relation  to  active  site  loop  dynamics.  Journal  of  Molecular  Biology  2001,  310  (1),  271-­‐280.    40.   Berlow,   R.   B.;   Igumenova,   T.   I.;   Loria,   J.   P.,   Value   of   a   hydrogen   bond   in  triosephosphate  isomerase  loop  motion.  Biochemistry  2007,  46  (20),  6001-­‐6010.    41.   Carpenter,   R.   A.;   Xiong,   J.;   Robbins,   J.   M.;   Ellis,   H.   R.,   Functional   Role   of   a  Conserved   Arginine   Residue   Located   on   a   Mobile   Loop   of   Alkanesulfonate  Monooxygenase.  Biochemistry  2011,  50  (29),  6469-­‐6477.    42.  Lipchock,  J.;  Loria,  J.  P.,  Millisecond  dynamics  in  the  allosteric  enzyme  imidazole  glycerol   phosphate   synthase   (IGPS)   from   Thermotoga   maritima.   Journal   of  Biomolecular  Nmr  2009,  45  (1-­‐2),  73-­‐84.    43.  Liang,  W.;  Ouyang,  S.;  Shaw,  N.;  Joachimiak,  A.;  Zhang,  R.;  Liu,  Z.-­‐J.,  Conversion  of  D-­‐ribulose  5-­‐phosphate  to  D-­‐xylulose  5-­‐phosphate:  new  insights  from  structural  and  biochemical  studies  on  human  RPE.  Faseb  Journal  25  (2),  497-­‐504.    44.   List,   F.;   Sterner,   R.;   Wilmanns,   M.,   Related   (beta   alpha)(8)-­‐Barrel   Proteins   in  Histidine   and   Tryptophan   Biosynthesis:   A   Paradigm   to   Study   Enzyme   Evolution.  Chembiochem  12  (10),  1487-­‐1494.    45.  Crawford,  I.  P.,  Evolution  of  a  Biosynthetic-­‐Pathway  -­‐  the  Tryptophan  Paradigm.  Annual  Review  of  Microbiology  1989,  43,  567-­‐600.    46.   Murzin,   A.   G.;   Brenner,   S.   E.;   Hubbard,   T.;   Chothia,   C.,   SCOP   -­‐   A   structural  classification  fo  proteins  database  for  the  investigation  of  sequences  and  structures.  Journal  of  Molecular  Biology  1995,  247  (4),  536-­‐540.    47.  Leopoldseder,  S.;  Claren,  J.;  Jurgens,  C.;  Sterner,  R.,  Interconverting  the  catalytic  activities   of   (beta   alpha)(8)-­‐barrel   enzymes   from   different   metabolic   pathways:  Sequence  requirements  and  molecular  analysis.  Journal  of  Molecular  Biology  2004,  337  (4),  871-­‐879.    48.  Lang,  D.;  Thoma,  R.;  Henn-­‐Sax,  M.;  Sterner,  R.;  Wilmanns,  M.,  Structural  evidence  for   evolution   of   the   beta/alpha   barrel   scaffold   by   gene   duplication   and   fusion.  Science  2000,  289  (5484),  1546-­‐1550.    49.   Horowitz,   N.   H.,   ON   THE   EVOLUTION   OF   BIOCHEMICAL   SYNTHESES.  Proceedings   of   the   National   Academy   of   Sciences   of   the   United   States   of   America  1945,  31  (6),  153-­‐157.    50.   Claren,   J.;   Malisi,   C.;   Hocker,   B.;   Sterner,   R.,   Establishing   wild-­‐type   levels   of  catalytic   activity   on   natural   and   artificial   (beta   alpha)(8)-­‐barrel   protein   scaffolds.  

 

 

27  

Proceedings   of   the   National   Academy   of   Sciences   of   the   United   States   of   America  2009,  106  (10),  3704-­‐3709.    51.  Saab-­‐Rincon,  G.;  Olvera,  L.;  Olvera,  M.;  Rudino-­‐Pinera,  E.;  Benites,  E.;  Soberon,  X.;  Moretti,  E.,  Evolutionary  Walk  between  (beta/alpha)(8)  Barrels:  Catalytic  Migration  from   Triosephosphate   Isomerase   to   Thiamin   Phosphate   Synthase.   Journal   of  Molecular  Biology  2011,  416  (2),  255-­‐270.    52.   Evran,   S.;   Telefoncu,   A.;   Sterner,   R.,   Directed   evolution   of   (B/a)(8)-­‐barrel  enzymes:   establishing   phosphoribosylanthranilate   isomerisation   activity   on   the  scaffold  of  the  tryptophan  synthase  -­‐subunit.  Protein  Engineering  Design  &  Selection  2012,  25  (6),  285-­‐293.    53.  Ochoa-­‐Leyva,  A.;  Soberon,  X.;  Sanchez,  F.;  Arguello,  M.;  Montero-­‐Moran,  G.;  Saab-­‐Rincon,   G.,   Protein   Design   through   Systematic   Catalytic   Loop   Exchange   in   the  (beta/alpha)(8)  Fold.  Journal  of  Molecular  Biology  2009,  387  (4),  949-­‐964.    54.  Barden,  T.  C.,   Indoles:  Industrial,  agricultural  and  over-­‐the-­‐counter  uses.  Topics  in  Hetercyclic  Chemistry  2011,  26,  31-­‐46.    55.  Shen,  H.;  Wang,  F.;  Zhang,  Y.;  Huang,  Q.;  Xu,  S.;  Hu,  H.;  Yue,  J.;  Wang,  H.,  A  novel  inhibitor   of   indole-­‐3-­‐glycerol   phosphate   synthase  with   activity   against  multidrug-­‐resistant  Mycobacterium  tuberculosis.  Febs  Journal  2009,  276  (1),  144-­‐154.    56.  Czekster,  C.  M.;  Neto,  B.  A.  D.;  Lapis,  A.  A.  M.;  Dupont,  J.;  Santos,  D.  S.;  Basso,  L.  A.,  Steady-­‐state  kinetics  of   indole-­‐3-­‐glycerol  phosphate  synthase  from  Mycobacterium  tuberculosis.  Archives  of  Biochemistry  and  Biophysics  2009,  486  (1),  19-­‐26.    57.   Smith,   D.   A.;   Parish,   T.;   Stoker,   N.   G.;   Bancroft,   G.   J.,   Characterization   of  auxotrophic  mutants  of  Mycobacterium  tuberculosis  and  their  potential  as  vaccine  candidates.  Infection  and  Immunity  2001,  69  (2),  1142-­‐1150.    58.  Sassetti,  C.  M.;  Boyd,  D.  H.;  Rubin,  E.  J.,  Genes  required  for  mycobacterial  growth  defined  by  high  density  mutagenesis.  Molecular  Microbiology  2003,  48  (1),  77-­‐84.  59.   Siddiqi,   M.   I.;   Kumar,   A.,   Review   of   knowledge   for   rational   design   and  identification   of   anti-­‐tubercular   compounds.   Expert   Opinion   on   Drug   Discovery  2009,  4  (10),  1005-­‐1015.    60.   Lu,  J.;  Yue,  J.;  Wu,  J.;  Luo,  R.;  Hu,  Z.;  Li,  J.;  Bai,  Y.;  Tang,  Z.;  Xian,  Q.;  Zhang,  X.;  Wang,   H.,   In   vitro   and   in   vivo   Activities   of   a   New   Lead   Compound   I2906   against  Mycobacterium  tuberculosis.  Pharmacology  2010,  85  (6),  365-­‐371.    61.   Li,  W.  F.;  Zhou,  X.  X.;  Lu,  P.,  Structural  features  of  thermozymes.  Biotechnology  Advances  2005,  23  (4),  271-­‐281.  

 

 

28  

62.  Unsworth,  L.  D.;  van  der  Oost,  J.;  Koutsopoulos,  S.,  Hyperthermophilic  enzymes  -­‐  stability,  activity  and   implementation  strategies   for  high   temperature  applications.  Febs  Journal  2007,  274  (16),  4044-­‐4056.    63.   Feller,   G.,   Protein   stability   and   enzyme   activity   at   extreme   biological  temperatures.  Journal  of  Physics-­Condensed  Matter  2010,  22  (32).    64.  Merz,  A.;  Yee,  M.  C.;  Szadkowski,  H.;  Pappenberger,  G.;  Crameri,  A.;  Stemmer,  W.  P.  C.;   Yanofsky,  C.;  Kirschner,  K.,   Improving   the   catalytic   activity  of   a   thermophilic  enzyme  at  low  temperatures.  Biochemistry  2000,  39  (5),  880-­‐889.    65.   Hennig,   M.;   Darimont,   B.;   Sterner,   R.;   Kirschner,   K.;   Jansonius,   J.   N.,   2.0   A  structure   of   indole-­‐3-­‐glycerol   phosphate   synthase   from   the   hyperthermophile  Sulfolobus  solfataricus:  possible  determinants  of  protein  stability.  Structure  1995,  3  (12),  1295-­‐306.    66.  Knochel,  T.  R.;  Hennig,  M.;  Merz,  A.;  Darimont,  B.;  Kirschner,  K.;   Jansonius,  J.  N.,  The   crystal   structure   of   indole-­‐3-­‐glycerol   phosphate   synthase   from   the  hyperthermophilic  archaeon  Sulfolobus  solfataricus  in  three  different  crystal  forms:  effects  of  ionic  strength.  J  Mol  Biol  1996,  262  (4),  502-­‐15.    67.   Knochel,   T.;   Pappenberger,   A.;   Jansonius,   J.   N.;   Kirschner,   K.,   The   crystal  structure  of  indoleglycerol-­‐phosphate  synthase  from  Thermotoga  maritima  -­‐  Kinetic  stabilization  by  salt  bridges.   Journal  of  Biological  Chemistry  2002,  277   (10),  8626-­‐8634.    68.   Bagautdinov,   B.;   Yutani,   K.,   Structure   of   indole-­‐3-­‐glycerol   phosphate   synthase  from   Thermus   thermophilus   HB8:   implications   for   thermal   stability.   Acta  Crystallographica  Section  D-­Biological  Crystallography  2011,  67,  1054-­‐1064.    69.   Gao,   L.;   Danno,   A.;   Fujii,   S.;   Fukuda,   W.;   Imanaka,   T.;   Fujiwara,   S.,   Indole-­‐3-­‐Glycerol-­‐Phosphate  Synthase  Is  Recognized  by  a  Cold-­‐Inducible  Group  II  Chaperonin  in   Thermococcus   kodakarensis.   Applied   and   Environmental   Microbiology   78   (11),  3806-­‐3815.    70.  Mazumder-­‐Shivakumar,  D.;  Bruice,  T.  C.,  Molecular  dynamics  studies  of  ground  state   and   intermediate   of   the   hyperthermophilic   indole-­‐3-­‐glycerol   phosphate  synthase.   Proceedings   of   the   National   Academy   of   Sciences   of   the   United   States   of  America  2004,  101  (40),  14379-­‐14384.    71.  Wolf-­‐Watz,  M.;  Thai,  V.;  Henzler-­‐Wildman,  K.;  Hadjipavlou,  G.;  Eisenmesser,  E.  Z.;  Kern,   D.,   Linkage   between   dynamics   and   catalysis   in   a   thermophilic-­‐mesophilic  enzyme  pair.  Nature  Structural  &  Molecular  Biology  2004,  11  (10),  945-­‐949.    72.   Vemparala,   S.;   Mehrotra,   S.;   Balaram,   H.,   Role   of   loop   dynamics   in   thermal  stability  of  mesophilic   and   thermophilic   adenylosuccinate   synthetase:  A  molecular  

 

 

29  

dynamics  and  normal  mode  analysis   study.  Biochimica  Et  Biophysica  Acta-­Proteins  and  Proteomics  1814  (5),  630-­‐637.    73.  Schlee,  S.;  Dietrich,  S.;  Kurcon,  T.;  Delaney,  P.;  Goodey,  N.  M.;  Sterner,  R.,  Kinetic  Mechanism   of   Indole-­‐3-­‐glycerol   Phosphate   Synthase.   Biochemistry   2012,   52   (1),  132-­‐142.    74.  Houlihan,  W.  J.,  Indoles.  Wiley-­‐Interscience:  New  York,  1972;  Vol.  2.  75.   Smith,   O.   H.;   Yanofsky,   C.,   1-­‐(ORTHO-­‐CARBOXYPHENYLAMINO)-­‐1-­‐DEOXYRIBULOSE   5-­‐PHOSPHATE,   A   NEW   INTERMEDIATE   IN   THE   BIOSYNTHESIS  OF  TRYPTOPHAN.  Journal  of  Biological  Chemistry  1960,  235  (7),  2051-­‐2057.    76.  Darimont,  B.;  Stehlin,  C.;  Szadkowski,  H.;  Kirschner,  K.,  Mutational  analysis  of  the  active   site   of   indoleglycerol   phosphate   synthase   from   Escherichia   coli.   Protein  Science  1998,  7  (5),  1221-­‐1232.    77.   Hennig,   M.;   Darimont,   B.   D.;   Jansonius,   J.   N.;   Kirschner,   K.,   The   catalytic  mechanism  of  indole-­‐3-­‐glycerol  phosphate  synthase:  crystal  structures  of  complexes  of  the  enzyme  from  Sulfolobus  solfataricus  with  substrate  analogue,  substrate,  and  product.  J  Mol  Biol  2002,  319  (3),  757-­‐66.    78.  Mazumder-­‐Shivakumar,   D.;   Kahn,   K.;   Bruice,   T.   C.,   Computational   study   of   the  ground   state   of   thermophilic   indole   glycerol   phosphate   synthase:   Structural  alterations   at   the   active   site   with   temperature.   Journal   of   the   American   Chemical  Society  2004,  126  (19),  5936-­‐5937.    79.   Shen,   H.   B.;   Xu,   F.;   Hu,   H.   R.;   Wang,   F.   F.;   Wu,   Q.;   Huang,   Q.;   Wang,   H.   H.,  Coevolving   residues   of   (beta/alpha)(8)-­‐barrel   proteins   play   roles   in   stabilizing  active   site   architecture   and   coordinating   protein   dynamics.   Journal   of   Structural  Biology  2008,  164  (3),  281-­‐292.    80.   Fodor,  A.  A.;  Aldrich,  R.  W.,  Influence  of  conservation  on  calculations  of  amino  acid   covariance   in   multiple   sequence   alignments.   Proteins-­Structure   Function   and  Bioinformatics  2004,  56  (2),  211-­‐221.    81.   Dekker,   J.   P.;   Fodor,   A.;   Aldrich,   R.   W.;   Yellen,   G.,   A   perturbation-­‐based  method   for   calculating   explicit   likelihood   of   evolutionary   co-­‐variance   in   multiple  sequence  alignments.  Bioinformatics  2004,  20  (10),  1565-­‐1572.    82.   Estabrook,  R.  A.;  Luo,  J.;  Purdy,  M.  M.;  Sharma,  V.;  Weakliem,  P.;  Bruice,  T.  C.;  Reich,  N.  O.,  Statistical  colevolution  analysis  and  molecular  dynamics:  Identification  of   amino   acid  pairs   essential   for   catalysis.  Proceedings   of   the  National   Academy   of  Sciences  of  the  United  States  of  America  2005,  102  (4),  994-­‐999.    

 

 

30  

Chapter 2

The Temperature Dependent Kinetic Mechanism of Thermophilic and

Mesophilic IGPS Enzymes

[This Chapter was adapted from the paper entitled “Differences in the catalytic

mechanism between mesophilic and thermophilic indole-3-glycerol phosphate synthase

enzymes at their adaptive temperatures” by Margot J. Zaccardi, Olga Mannweiler, and

David D. Boehr in Biochemical and Biochemical Research Communications, 2012, 418,

324-329. Olga Mannweiler performed experiments on the IGPS enzyme from E. coli. All

other experiments were performed by Margot J. Zaccardi]1

2.1 Abstract

Thermophilic enzymes tend to be less catalytically-active at lower temperatures

relative to their mesophilic counterparts, despite having very similar crystal structures.

An often cited hypothesis for this general observation is that thermostable enzymes have

evolved a more rigid tertiary structure in order to cope with their more extreme, natural

environment, but they are also less flexible at lower temperatures, leading to their lower

catalytic activity under mesophilic conditions. An alternative hypothesis is that

complementary thermophilic–mesophilic enzyme pairs simply operate through different

evolutionary-optimized catalytic mechanisms. In this Chapter, we present evidence that

while the steps of the catalytic mechanisms for mesophilic and thermophilic indole-3-

glycerol phosphate synthase (IGPS) enzymes are fundamentally similar, the identity of

the rate-determining step changes as a function of temperature. Our findings indicate that

 

 

31  

while product release is rate-determining at 25 °C for thermophilic IGPS, near its

adaptive temperature (75 °C), a proton transfer event, involving a general acid, becomes

rate-determining. The rate-determining steps for thermophilic and mesophilic IGPS

enzymes are also different at their respective, adaptive temperatures.

2.2 Introduction

Investigations into biological temperature adaptation and enzyme stability at

extreme temperatures can provide a deeper understanding of life under extreme

conditions, and can aid in the design of enzymes for industrial applications and new

biocatalysts with a wide range of temperature optima.2, 3 The use of thermophilic

enzymes for industrial processes provides the opportunity to improve a variety of

common synthesis issues including solubility, reaction time, and product yield.

Additionally, reactions performed at temperatures nearing 100 °C considerably decreases

the risk of bacterial contamination for food and drug related biosyntheses.4

High temperature increases the fluidity of membranes and destroys the normal

activity of biomolecules including proteins. Thermophilic organisms have evolved a

range of mechanisms to combat these problems in order to sustain life at higher

temeperatures.2 To maintain their three-dimensional structures at higher temperatures,

thermophilic enzymes tend to have an increased number of noncovalent interactions, such

as salt bridges and/or disulfide bonds, when compared to their mesophilic homologs.

Despite the similarities in amino acid sequences and three-dimensional structures, warm-

adapted enzymes have lower catalytic activity than their mesophilic counterparts when

assayed at lower temperatures.5, 6 The additional interactions required for stability are

 

 

32  

thought to limit enzyme flexibility at lower temperatures, leading to a reduction in

enzyme activity.5-7 However, thermostable enzymes can still contain flexible regions that

are important or required for function as was seen by Wolf-Watz et al. in a comparison of

adenylate kinase from E. coli and the hyperthermophile Aquifex aeolicus.8

The purpose of this study is to investigate the differences in the kinetic

mechanism for IGPS enzymes from thermophilic versus mesophilic organisms, and to

determine whether other factors besides protein flexibility must be considered when

analyzing this temperature dependent activity. A more detailed explanation would

involve not only changes in enzyme flexibility, but also variations in the catalytic

mechanism due to adaptations required for structural stability and activity at higher

temperatures. Thermophilic enzymes from the tryptophan biosynthetic pathway,

including IGPS from Thermus thermophilis, T. maritima (tmIGPS), and S. sulfataricus

(ssIGPS) have been used as model systems to decipher the interactions responsible for

protein thermostability.9-12 S. sulfataricus, the thermophile of interest in these studies, is

found in hot sulfur beds,2 and ssIGPS is stable at temperatures above 85 °C.9

Despite a sequence identity of only 30%, ssIGPS and its mesophilic counterpart

from Escherichia coli (ecIGPS) show a strong structural similarity with a root mean

square deviaton (rmsd) of only 1.73 Å (Figure 2.1).9 The high thermostability of ssIGPS

is largely attributed to the increased occurrence of salt bridges (ssIGPS contains thirteen

additional salt bridges compared to ecIGPS) that connect loops and helices, creating a

tight network within the enzyme. The maximum catalytic turnover rate constant (kcat) for

ssIGPS is much lower at 37 °C compared to ecIGPS. The difference in activity was

 

 

33  

Figure 2.1: Conserved structure and function of IGPS from E. coli (green) (PDB 1P11) and S. sulfobolus (blue) (PDB 1IGS). Despite only 30% sequence identity and large differences in stability, ssIGPS and ecIGPS show strong structural similarity.  

 

 

34  

previously attributed to the lower flexibility of ssIGPS at this temperature.9, 11 Previous

studies suggested that the rate-determining step of ssIGPS at 25 °C is product release.13

However, this temperature is not necessarily relevant for understanding the biological

activity of the thermophilic enzyme. We propose that while the chemical mechanisms for

thermophilic and mesophilic IGPS are similar, the kinetic parameters governing these

processes have different temperature dependencies. In this Chapter, we have examined

the kinetic mechanism for ssIGPS and ecIGPS over a range of temperatures. The results

show that the rate-determining step for ssIGPS is temperature dependent. While product

release is rate-determining at 25 °C, at higher, biologically relevant temperatures (i.e. 75

°C), the ring closure step of the chemical reaction is rate-determining. Additionally, the

rate-determining step for ecIGPS at its adaptive temperature (37 °C) is different from

ssIGPS.

2.3 Experimental Methods

2.3.1 Cloning of ssIGPS and ecIGPS

In E. coli, IGPS is found as the N-terminal domain in a bifunctional enzyme

covalently linked to the previous enzyme in the biosynthetic pathway, PRAI. IGPS in S.

sulfataricus occurs as a monofunctional enzyme. The ecIGPS monofunctional domain

has been shown to retain catalytic ability that is comparable to the bifunctional

complex;14 for the studies presented herein, only the ecIGPS domain was used (amino

acids 1–259). The E. coli codon usage-optimized transcript for both ssIGPS and ecIGPS

(GenScript) were PCR-amplified from the plasmid pSC101-trp (ATCC 31743), and

cloned into pET101 (Ampicillin, AmpR) using the Champion pET Directional TOPO

 

 

35  

Expression Kit (Invitrogen). Unfortunately, preliminary protein expression trials

indicated a substantial overexpression of the β-lactamase protein. Consequently, the

genes were sub-cloned from pET101 into pET26 (Kanamycin, KanR) with the restriction

endonucleases XbaI and SacI using standard procedures. Due to low overexpression of

ssIGPS using the pET26 construct, the ssIGPS gene in pET21b (AmpR) was also obtained

as a kind gift from Dr. Reinhard Sterner at Universitaet Regensburg (Germany) along

with tmIGPS in the pET21b construct. The Lys110Arg ssIGPS variant was generated

using the QuikChange Lightning kit® (Stratagene) with appropriate primers for the E. coli

optimized pET26 ssIGPS construct. All sequences for wild type (WT) and amino acid

substituted IGPS were verified through DNA sequencing (Nucleic Acid Facility, The

Pennsylvania State University).

2.3.2 Overexpression and Purification of ssIGPS, ecIGPS, and tmIGPS

The overexpression of ssIGPS and ecIGPS was performed by transforming

plasmids (pET26) carrying the genes into E. coli BL21(DE3)star cells. A 10 mL Luria-

Bertani (LB) starter culture was inoculated with fresh transformations on LB-Agar

containing 50 µg/mL Kan and grown overnight at 37 °C. This culture was subsequently

used to inoculate 1L of LB media. The culture was grown at 37 °C to an optical density

(A600) between 0.500 and 0.600 at which time it was induced with isopropyl β-D-1-

thiogalactopyranoside (IPTG) and allowed to grow for approximately 20 hours at 25°C.

Cells were harvested by centrifugation (10,000 x g, 4°C, 20 minutes).

The purification of the enzymes followed protocols similar to those previously

described.9, 15 For ssIGPS, cell pellets were resuspended in 30 mL of 100 mM potassium

 

 

36  

phosphate pH 7.8, 2 mM ethylenediamine tetraacetic acid (EDTA) and 1 mM

phenylmethanesulfonylfluoride (PMSF), and were lysed through sonication. Cell lysates

were then centrifuged at 30,000 xg and 4 °C for 30 min. The supernatant was heated at 75

°C for ten minutes to precipitate thermolabile host proteins, which were separated out via

centrifugation. The supernatant was then dialysed against 10 mM potassium phosphate,

pH 7.8, 2 mM EDTA (buffer A) and applied to a 20 mL HiPrep 16/10 Q-Sepharose anion

exchange column (GE Healthcare), washed with buffer A and eluted using a phosphate

gradient (0–100 mM phosphate over 20 column volumes, flow rate = 2 mL/min).

Fractions containing ssIGPS were selected using SDS–PAGE (sodium dodecylsulfate-

polyacrylamide electrophoresis), pooled and concentrated to approximately 1 mL using

Vivaspin-20 spin concentrators (Sartorius Stedium Biotech). Partially purified ssIGPS

was applied to a HiPrep 16/60 Sepharcryl S100 gel filtration column (GE Healthcare) and

eluted using buffer A containing 200 mM NaCl. Fractions containing ssIGPS were again

selected via SDS-PAGE, pooled, concentrated and then dialyzed against buffer A. The

protocol for ecIGPS was similar except that the cell pellet was reconstituted directly into

the Q-sepharose buffer A (50 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid

(HEPES) pH 7.5, 1 mM EDTA) and eluted on the anion exchange column with a 0 to

500M gradient of NaCl.

The pET21b ssIGPS construct is hexa-histidine tagged (His-tagged); the

overexpression and purification differed from that described for the pET26 construct.16

The growth was performed with BL21-CodonPlus(DE3)-RIPL (Agilent) E. coli cells in

LB media containing 100 µg/mL Amp following the same protocol as described for the

pET26 ssIGPS construct. These cells are advantageous because they are optimized for the

 

 

37  

overexpression of heterologous proteins in E. coli. The cell pellet was reconstituted in 30

mL 100mM potassium phosphate pH 7.0, 300 mM potassium chloride (KCl) (lysis

buffer), and 1 mM PMSF, and were lysed through sonification. Cell lysates were then

centrifuged at 30,000 xg and 4 °C for 30 min, and the supertenant was heated at 75 °C for

ten minutes to precipitate thermolabile contaminants, which were removed via

centrifugation. The subsequent supernatant was applied to a column containing 2mL Ni-

NTA (HisPur-ThermoScientific). The column was then washed with lysis buffer plus 10,

20, 50, 100, and 250 mM Imidazole to elute bound protein. Fractions were selected by

SDS-PAGE, pooled, and concentrated as described.

The overexpression and purification protocols for tmIGPS, which is also in the

pET21b construct with a hexa-histidine tag, were identical to that for hexa-his tagged

ssIGPS except that no PMSF was added to the lysis buffer as described previously by

Schneider and coworkers.16

2.3.3 Steady-State Kinetic Assays for IGPS

ssIGPS activity was measured as previously described via fluorescence by

monitoring the formation of IGP.1, 14, 17 The sample was excited at 278 nm and emission

measured at 340 nm using a spectrofluorometer (Horiba Jobin Yvon). Assays were

performed in 50 mM 3-[4-(2-hydroxyethyl)-1-piperazinyl] propanesulfonic acid (HEPPS)

pH 7.5, 4 mM EDTA unless otherwise noted with an enzyme concentration of 10 nM and

a volume of 300 µL with substrate concentrations ranging from 30 nM to 4 µM. Raw

rates in cps/s were converted to nM/s using a standard curve for cps versus concentration

of CdRP (Figure 2.2). This standard curve was obtained using tmIGPS since that enzyme

 

 

38  

Figure 2.2: Standard curve of fluorescence units per nanomolar for converting cps/s to nM/s. The slope of the line (4036 cps/nM) was used to convert data for ssIGPS to the appropriate units. Curve was attained using IGPS from T. maritima, which does not display product inhibition.

 

 

39  

does not exhibit product inhibition.16 Assay times ranged from 30 seconds to 60 seconds.

ecIGPS activity was measured by monitoring the formation of IGP via absorbance

at 278 nm on a SpectraMax M2 plate reader (Molecular Devices). Assays were

performed in 50 mM HEPPS pH 7.5, 4 mM EDTA using a 96-well microtiter plate assay.

Assay volume was 250 µL, substrate concentrations ranged from 100 nM to 60 µM, and

assays were typically conducted for three to four minutes. Initial rates were fit to the

Michaelis–Menten equation (Equation 2.1) using nonlinear regression with

Kaleidograph,

€

v =kcat[E]T [S]KM + [S] (2.1)

where ν is the initial velocity, ET is the total enzyme concentration, and [S] is the

concentration of substrate, CdRP. All assays were performed in triplicate. Representative

progress curves and Michaelis-Menton curves for ssIGPS are shown in Figure 2.3.

2.3.4 Solvent Viscosity Effects, Solvent Deuterium Kinetic Isotope Effects, and pH

Effects

The solvent viscosity effects (SVE), solvent deuterium kinetic isotope effects

(SDKIE) and pH effects for IGPS enzymes were determined by varying the buffer

conditions of the standard assay. SVEs were determined using enzyme assays in 50 mM

HEPPS pH 7.5, 4 mM EDTA buffer with 0 to 30 % (w/v) glycerol. SVE experiments

were also performed in an alternate viscogen, sucrose, as well as in a microviscogen,

PEG 8000, as controls for nonspecific viscogen effects. Assays were performed at

saturating substrate concentrations (800 µM for ssIGPS; 12 µM for ecIGPS). The relative

viscosities of the buffer solutions were measured using an Ostwald viscometer. The SVE

 

 

40  

Figure 2.3: Representative data for ssIGPS assays. (a) Progress curves for ssIGPS at 75 °C at varying concentrations of CdRP (100, 400, 800, 1000, and 2000 nM). (b) Michaelis-Menton curve for ssIGPS at 75 °C.

 

 

41  

is defined as the slope of the line for the plot of (ratewithout viscogen/ratewith viscogen) versus

relative viscosity.

SDKIEs were obtained by comparing IGPS enzyme activities in H2O and D2O.

The pD was used instead of pH for solutions in D2O and was adjusted according to pD =

pH + 0.4. Proton inventory studies were performed by varying the mole fraction of D2O

in the buffer from 0 to 1.

pH studies for ssIGPS (37 and 75 °C) and ecIGPS (37 °C) were performed in

buffers with overlapping buffering ranges, including: 100 mM 2-(N-morpholino) ethane

sulfonic acid (MES) (pH 5.0-6.5), 100 mM HEPES (pH 6.5-8.0), N,N-Bis(2-

hydroxyethyl) glycine (BICINE) (pH 8.0-8.5), and 100 mM N-cyclohexyl-2-

aminoethanesulfonic acid (CHES) (pH 8.6-10.0). The pH rate profiles for both ecIGPS

and ssIGPS display two ionizations, and were fit to Eq. (2) using Kaleidograph:

(2.2)

where ν is the estimated kcat, C is the pH-independent rate value, and pKa1 and pKa2 are

the pKa values associated with the ascending and descending limbs of the pH profile,

respectively.

2.3.5 Synthesis of CdRP

Originally, the protocol described by Czekster et al.18 was used to synthesize the

solid barium salt of the substrate, CdRP. However, preliminary assays showed that this

form of the substrate was not active in assays measuring for IGPS activity and a new

protocol was obtained.15

!

" = C /(1+10pKa1# pH +10pH# pKa 2 )

 

 

42  

In short, 47.7 mg of anthranilic acid was dissolved in 200 µL of ethanol and 58.8

mg of ribose-5-phosphate was dissolved in 200 µL of water. The two solutions were

combined and allowed to stand at room temperature in the dark for twenty hours. After

incubation, 4 mL of water was added to the reaction mixture and allowed to stand in the

dark for one hour. The product was washed with 5 mL ethyl acetate, allowing separation

of the aqueous and solvent layers in the separatory funnel. The aqueous phase containing

the CdRP was separated and the wash with ethyl acetate was repeated approximately five

to seven times until the solvent phase was colorless. The product was then purged with

nitrogen to remove any remaining ethyl acetate and stored in the dark at -80°C.

The concentration of CdRP was calculated using the tmGPS enzyme to convert

CdRP to IGP and measuring using absorbance both the decrease in CdRP at 327 nm

(ε=3.43 mM-1cm-1) and the increase in IGP at 278 nm (ε=4.48 mM-1cm-1).

2.3.6 Circular Dichroism

Circular dicroism (CD) experiments were performed on WT and Lys110Arg

ssIGPS on a Jasco J-810 Spectropolarimeter from 250 nm to 190 nm with 1 nm intervals

and a 1 nm bandwidth. The experiments were performed in 10 mM potassium phosphate

pH 7.0 with an enzyme concentration of 1.7 µM.

 

 

43  

2.4 Results

2.4.1 Steady-state Kinetic of ssIGPS

The kinetic mechanism for IGPS can be described as follows:19

€

E + S⇔k−1

k1ES⇔

k−2

k2ES *⇔

k−3

k3EI1→

k4EI2⇔

k−5

k5EP *⇔

k−6

k6EP⇔

k−7

k7E + P

(2.3)

In this mechanism, the substrate (S = CdRP) binds to the enzyme to form the enzyme-

substrate complex (ES), which may then undergo a conformational change to form the

more active ES* complex, followed by irreversible chemistry involving the release of

CO2 gas to yield the enzyme-product (P = IGP) complex. This step likely reflects the

chemistry (condensation, decarboxylation, dehydration) proposed for IGPS, which occurs

through two intermediates, I1 and I2. The product complex then must undergo another

conformational change to allow product release in the final step.

To comprehensively compare the catalytic activities of ssIGPS and ecIGPS, the

steady-state kinetic parameters of the IGPS enzymes were determined across multiple

temperatures (25, 37, and 75 °C), ensuring that biologically relevant data was obtained

for both enzymes (Table 2.1). Consistent with previous observations, the maximum

turnover rate (kcat) for ecIGPS was 20-fold greater than that for ssIGPS at both 25°C and

37 °C, although the enzymes were similar in their catalytic efficiencies (kcat/KM).11 The

binding affinity of the substrate for ssIGPS is temperature independent, with similar KM

values at all temperatures assayed (~50 to 100 nM). Conversely, kcat decreases as

temperature decreases (0.67 s-1 at 75 °C versus 0.16 s-1 at 25 °C). The previously

proposed rate-determining step of ssIGPS at 25 °C is the release of product from the

enzyme,19 but the rate-determining step at more higher temperatures is not known.

 

 

44  

Table 2.1: Steady-state kinetic parameters for ssIGPS and ecIGPS at pH 7.5 indicate that the rate-determining step changes as a function of temperature. Sample Temp

(°C) kcat (s-1) KM (nM) kcat/KM (x

106 M-1s -1) SVE SDKIE

ssIGPS 25 0.16 ± 0.02

74 ± 38 2.2 1.0 ± 0.2a 1.2 ± 0.2a

37 0.42 ± 0.04

88 ± 47 4.8 0.6 ± 0.3a 5.8 ± 0.1a

75 0.67 ± 0.03

44 ± 9 15 -0.2 ± 0.1a 3.6 ± 0.3a

ecIGPSc 25 4.1 ± 0.2a n.d.b n.d.b n.d.b n.d.b

37 9.3 ±0.6 1600 ± 300

5.7 0.2 ± 0.1a 1.0 ± 0.1a

aValues for kcat were determined using saturating substrate concentrations (800 nM for ssIGPS and 12 µM for ecIGPS) bValues not determined. cAssays on ecIGPS were performed by Olga Mannweiler

 

 

45  

2.4.2 Solvent Viscosity Effects, Solvent Deuterium Kinetic Isotope Effects, and pH

Effects

There are various limiting scenarios for the kinetic mechanism of ssIGPS that will

determine the value of kcat : if product release is relatively fast compared to the chemical

step(s) (i.e. k3 << k5), then kcat will report on k3. Conversely, if product release is rate-

determining (i.e. k3 >> k5), then kcat will approach k5. These scenarios can be resolved

based on the sensitivity of the turnover rate to increasing solvent viscogen and the

introduction of deuterated solvent. Diffusion-limited processes including substrate

binding, product release, or large, global conformational changes will be dependent on

the viscosity of the solution, whereas processes independent of diffusion, like the

chemical steps, will be unaffected. Conversely, chemical processes whose rates are

dependent on the transfer of a solvent exchangeable proton will be affected by the

introduction of deuterated solvent as in the SDKIE experiments (Figure 2.2). In the IGPS

enzyme, the ring closure step is predicted to be isotope sensitive, due to the proton

donation from the general acid. Conversely, the dehydration the dehydration step is not

expected to produce a substantial SDKIE. Within the dehydration step, it is likely that the

removal of the non-exchangeable alkyl hydrogen is rate-determining over the loss of

water, causing the step to be isotope insensitive.

Solvent viscosity effects (SVEs) are defined by the slope of a plot of relative rate

(ratewithout viscogen/ratewith viscogen) versus relative viscosity, for which the theoretical

maximum effect is one. SVEs were determined for ecIGPS and ssIGPS enzymes across

multiple temperatures (Table 2.1, Figure 2.3). For ecIGPS, there is only a small SVE

(~0.20) at 37 °C. In contrast, there was a much larger SVE for ssIGPS at both 25 °C and

 

 

46  

Figure 2.4: The rate-determining step of the IGPS reaction can be deciphered using SVE and SDKIE experiments. Substrate binding and product release (green) are viscosity sensitive and isotope insensitive. Ring closure (blue) is viscosity insensitive isotope sensitive. Decarboxylation and dehydration are both viscosity and isotope insensitive.

 

 

47  

Figure 2.5: Solvent viscosity effects for ssIGPS. At 25 °C (blue) there is an SVE of 1.0 ± 0.2, wherease at 75 °C (black) the SVE is no longer present (-0.2 ± 0.1). The SVE is defined by the slop of the line for vo/vi versus ni/no. The results indicate that at 25 °C product release is rate-determining but as temperature increases to 75 °C product release is no longer rate-determining, and a chemical step becomes rate-determining.  

 

 

48  

37 °C, consistent with previous data suggesting that product release is rate-determining

for ssIGPS at these temperatures. However, there were not appreciable SVEs for ssIGPS

at higher temperatures (75 °C), reflecting a change in the identity of the rate-determining

step with increasing temperatures. The similar SVEs for ecIGPS and ssIGPS at their

respective, adaptive temperatures may indicate similar rate-determining step(s).

To test for any nonspecific effects of due to the introduction the microviscogen

used in these experiments (glycerol), the SVE experiments were also performed in

sucrose. The two different viscogens showed similar effects. At 37 °C, an SVE of 0.6 was

measured in glycerol for ssIGPS, and one of 0.4 was measured in sucrose. Similarly at 75

°C, an SVE of -0.2 was measured in both viscogens. This result indicates that the

viscosity effect is reporting on the diffusion controlled processes of IGPS catalysis.

Additionally, the experiments were performed in the presence of a macroviscogen, PEG

8000, in which the enzyme does not exhibit a viscosity effect (i.e. kcat(PEG 8000)/kcat(no

viscogen) was equal to 1.0 ± 0.2 for 1.9% (w/v) PEG 8000 at a relative viscosity of 1.7).

Differences in enzyme activities for ecIGPS and ssIGPS may be due to variations

in their catalytic mechanisms. The proposed mechanism for ssIGPS suggests the

involvement of both a general acid and a general base; if the chemical mechanism is the

same for ecIGPS and ssIGPS, a similar pH rate profile should be observed. The pH

dependence of the activity for ecIGPS and ssIGPS was determined (Figure 2.4, Table

2.2). The pH rate profiles for ecIGPS and ssIGPS at biologically relevant temperatures

(37 °C for ecIGPS and 75 °C for ssIGPS) exhibit a bell shaped curve with both ascending

and descending limbs, consistent with the involvement of general base and acid

chemistry, respectively. These results reflect the pH dependence of kcat. The pH rate

 

 

49  

Figure 2.6: The pH dependence of WT ssIGPS at (a) 37 °C (pKa1 7.5 ± 0.2, pKa2 8.8 ± 0.3) and (b) 75 °C (pKa1 5.6 ± 0.2, pKa2 8.7 ± 0.2) and (c) ecIGPS at 37 °C (pKa1 6.7 ± 0.1, pKa2 8.8 ± 0.1) show an ascending and descending limb consistent with general base and general acid involvement, respectively.  

 

 

50  

Table 2.2: pKa values for ssIGPS and ecIGPS. Enzyme Temperature pKa1 pKa2

ecIGPS 37 °C 6.7 ± 0.1 8.8 ± 0.3

ssIGPS 37 °C 7.5 ± 0.2 8.8 ± 0.3

ssIGPS 75 °C 5.6 ± 0.2 8.7 ± 0.2

 

 

51  

profile of kcat/KM for ssIGPS at 75 °C also yielded a bell-shaped curve with similar pKa

values (5.0 ± 0.4, 9.3 ± 0.5). It should be noted that for IGPS from M. tuberculosis

(mtIGPS), only the ascending limb in the pH rate profile is observed with a pKa of 6.8,18

which is similar to the pKa for ecIGPS (6.70 ± 0.08). At 75 °C, the pKa value (5.6 ± 0.2)

for the ascending limb of ssIGPS is 1 pH unit lower compared to the mtIGPS and ecIGPS

enzymes. The descending limb for the pH rate profiles yielded similar pKas for ecIGPS

(8.7 ± 0.1) and ssIGPS (37 °C, 8.8 ± 0.3; 75 °C, 8.7 ± 0.2).

The pKa for the descending limb of both ecIGPS and ssIGPS is in the range

expected for a Lys residue, such as the proposed general acid, Lys110. To further test the

importance of Lys110, the Lys110Arg substitution was assayed. An Arg at this position

is expected to make similar noncovalent interactions (e.g. hydrogen bond, electrostatic

interactions) with the CdRP, but would act as a much less effective general acid with a

higher pKa (~12). Consistent with this suggestion, the Lys110Arg variant of ssIGPS had

very low enzyme activity (kobs < 1.5 x 10-5 s-1 at 75 °C) compared to WT ssIGPS (kcat =

0.67 ± 0.03 s-1 at 75 °C). Circular dichroism experiments were performed on both WT

and Lys110Arg ssIGPS to ensure that the variant was properly folded and the loss in

activity was not due to gross changes in secondary structure.

The IGPS-catalyzed reaction is proposed to proceed through three chemical steps.

SDKIEs offer a way to determine which of the chemical steps may be rate-determining

for the IGPS enzymes. For ssIGPS at 25 °C, there was not a significant SDKIE for kcat

(Table 2.1). This finding is consistent with the large SVEs and previous studies

suggesting that product release is fully rate-determining at this temperature.13, 19

However, at higher temperatures there was a substantial SDKIE for kcat (kH2O/kD2O = 3.6

 

 

52  

± 0.3 for ssIGPS at 75 °C). In contrast, ecIGPS does not have a substantial SDKIE for kcat

at 37 °C, suggesting that an isotope insensitive chemical step (and/or a viscosity-

independent conformational change) must be rate-determining. Consistent with the

ecIGPS results, mtIGPS also displayed a substantially lower SDKIE for kcat (~1.6)18

compared to ssIGPS. Parry’s proposed chemical mechanism (Figure 1.2) and the pH rate

profiles (Figure 2.4) for the IGPS enzymes suggest multiple proton transfer events that

may be responsible for the SDKIE observed for ssIGPS. To identify the responsible

proton transfer event(s) that are being reported on by the SDKIE experiment, proton

inventory studies and the pH dependence of the SDKIE were determined. At both 37 and

75 °C, the proton inventory study yielded a linear relationship between the relative rate

and mole fraction of D2O (Figure 2.5), indicating that a single proton transfer event is

responsible for the observed SDKIE for kcat. Additionally, for ssIGPS at pH 8.5, there

was not a substantial SDKIE for kcat (0.99 ± 0.07) at 37 °C, whereas at pH 6.5, an SDKIE

is still observed (2.18 ± 0.06), indicating that the SDKIE likely arises from the general

acid, Lys110.

 

 

53  

Figure 2.7: The rate-determining step for ssIGPS at higher temperatures involves a single proton transfer event. (a) The maximum catalytic turnover of ki/ko versus mole fraction D2O:H2O at both 37 °C (blue) and 75 °C (green) show a linear fit. (b) The square root of ki/ko versus mole fraction D2O:H2O at 37 °C and 75 °C show a quadratic fit. These results indicate that one proton transfer event is involved in the rate-determining step of the reaction, namely the proton transfer from the general acid in the condensation step of the reaction.  

 

 

54  

2.5 Discussion

2.5.1 Temperature Dependent Kinetic Mechanism of ssIGPS

Thermophilic enzymes show lower activity at lower temperatures compared to

higher temperatures and as compared to their mesophilic counterparts. This finding is

often attributed to changes in the flexibility of the enzyme at lower temperatures. In the

case of ssIGPS, while flexibility may have a role in the activity of the enzyme, the results

also indicate that changes in the rate-determining step of the reaction are largely

responsible for the apparent changes in activity. At its biologically relevant temperature

(75 °C), ssIGPS does not exhibit an SVE. The substantial SDKIE indicates that chemistry

is rate-determining. These results, along with the proton inventory study and the studies

on Lys110Arg, suggest that a single proton transfer event involving the general acid,

Lys110, is rate-determining for ssIGPS at higher temperatures. This result suggests that

the identity of the rate-determining step of the ssIGPS reaction at biologically relevant

temperatures is the initial ring closure step. As temperature decreases, the rate-

determining step changes from chemistry to product release, as evidenced by the

presence of a SVE, and which is consistent with previously published work.11

At 37 °C, both an SVE and SDKIE are present, indicating that multiple steps are

contributing to the kinetic parameters at this temperature; product release and chemistry

are both partially rate-determining at this temperature. The pH rate profile at 37 °C still

shows both ascending and descending limbs that are indicative of acid base catalysis, but

the trend is more complex, particularly for the ascending limb. In mtIGPS, the ascending

limb was previously attributed to the general base (Glu159 according to ssIGPS

numbering). In ssIGPS, at 37 °C, the poor fit to the curve in this  region (Figure 2.3) may

 

 

55  

be explained by allowing more than one responsible ionizable group, which would be

consistent with multiple kinetic steps contributing to the overall rate. This finding

suggests that a different ionizable group may be responsible for this pKa at different

temperatures, and/or different microenvironments within the IGPS active site may have

different effects on similar ionizable group(s). The lower pKa observed for ssIGPS at 75

°C is closer to the range expected for a Glu or Asp residue. This is consistent with the

proposed mechanism in which Glu159 or Glu210 acts as the general base in the

dehydration step.

2.5.2 Differences in the Rate-Determining Step of Thermophilic ssIGPS and

Mesophilic ecIGPS

While IGPS from both thermophilic and mesophilic organisms is believed to

undergo the same chemical steps toward the product, the enzyme takes may have evolved

differently due to the environmental restrictions for each organism. These changes can

manifest as changes in the kinetic mechanism and the rate-determining step of the

reaction. In the ecIGPS enzyme, only a small SVE and no SDKIE is seen at its

biologically active temperature, 37 °C, suggesting that a different chemical step such as

dehydration which does not involve a proton transfer, or a viscosity-independent

conformational  change, is largely rate-determining compared to ssIGPS. This result also

indicates differences in the rate-determining step for mesophilic and thermophilic IGPS

enzymes at their respective, adaptive temperatures.

The active site of IGPS enzymes are well shielded from solvent;9, 20 therefore,

conformational changes must accompany product release. At lower temperatures, these

 

 

56  

conformational changes are slow and product release is rate-determining for ssIGPS, and

at higher temperatures, the protein motions are sufficiently fast to relieve this bottleneck

to ssIGPS catalysis. However, even at their respective, adaptive temperatures, the rate-

determining steps for ssIGPS and ecIGPS differ. It is unlikely that the fundamental

chemical mechanisms are substantially different considering the amino acid sequence and

structural similarity of the IGPS enzymes.

2.6 Conclusions

In this chapter, the kinetic mechanism of IGPS was examined in both

thermophilic and mesophilic enzymes. The results suggest that IGPS enzymes have

evolved to more efficiently catalyze different steps of the chemical reaction(s), likely due

to the different environmental stressors present. Engineering thermostable enzymes or

new enzyme activities on thermophilic proteins like ssIGPS21, 22 will require careful

consideration of not only protein stability-flexibility relationships, but also a thorough

understanding of how different physical and chemical barriers to catalysis respond to

temperature.

2.7 References

1.   Zaccardi,   M.   J.;   Mannweiler,   O.;   Boehr,   D.   D.,   Differences   in   the   catalytic  mechanisms  of  mesophilic   and   thermophilic   indole-­‐3-­‐glycerol   phosphate   synthase  enzymes  at  their  adaptive  temperatures.  BBRC  2012,  418  (2),  324-­‐329.    2.  Rothschild,  L.  J.;  Mancinelli,  R.  L.,  Life  in  extreme  environments.  Nature  2001,  409  (6823),  1092-­‐1101.    3.  Egorova,  K.;  Antranikian,  G.,  Industrial  relevance  of  thermophilic  Archaea.  Current  Opinion  in  Microbiology  2005,  8  (6),  649-­‐655.    

 

 

57  

4.  Unsworth,  L.  D.;  van  der  Oost,   J.;  Koutsopoulos,  S.,  Hyperthermophilic  enzymes  -­‐  stability,  activity  and   implementation  strategies   for  high   temperature  applications.  Febs  Journal  2007,  274  (16),  4044-­‐4056.    5.   Feller,   G.,   Protein   stability   and   enzyme   activity   at   extreme   biological  temperatures.  Journal  of  Physics-­Condensed  Matter  2010,  22  (32).    6.  Sterpone,  F.;  Melchionna,  S.,  Thermophilic  proteins:  insight  and  perspective  from  in  silico  experiments.  Chemical  Society  Reviews  41  (5),  1665-­‐1676.    7.   Li,  W.   F.;   Zhou,   X.   X.;   Lu,   P.,   Structural   features   of   thermozymes.  Biotechnology  Advances  2005,  23  (4),  271-­‐281.    8.  Wolf-­‐Watz,  M.;  Thai,  V.;  Henzler-­‐Wildman,  K.;  Hadjipavlou,  G.;  Eisenmesser,  E.  Z.;  Kern,   D.,   Linkage   between   dynamics   and   catalysis   in   a   thermophilic-­‐mesophilic  enzyme  pair.  Nature  Structural  &  Molecular  Biology  2004,  11  (10),  945-­‐949.    9.  Hennig,  M.;  Darimont,  B.;  Sterner,  R.;  Kirschner,  K.;  Jansonius,  J.  N.,  2.0  A  structure  of   indole-­‐3-­‐glycerol   phosphate   synthase   from   the   hyperthermophile   Sulfolobus  solfataricus:   possible   determinants   of   protein   stability.   Structure   1995,   3   (12),  1295-­‐306.    10.  Knochel,  T.  R.;  Hennig,  M.;  Merz,  A.;  Darimont,  B.;  Kirschner,  K.;   Jansonius,  J.  N.,  The   crystal   structure   of   indole-­‐3-­‐glycerol   phosphate   synthase   from   the  hyperthermophilic  archaeon  Sulfolobus  solfataricus  in  three  different  crystal  forms:  effects  of  ionic  strength.  J  Mol  Biol  1996,  262  (4),  502-­‐15.    11.  Merz,  A.;  Yee,  M.  C.;  Szadkowski,  H.;  Pappenberger,  G.;  Crameri,  A.;  Stemmer,  W.  P.  C.;   Yanofsky,  C.;  Kirschner,  K.,   Improving   the   catalytic   activity  of   a   thermophilic  enzyme  at  low  temperatures.  Biochemistry  2000,  39  (5),  880-­‐889.    12.   Bagautdinov,   B.;   Yutani,   K.,   Structure   of   indole-­‐3-­‐glycerol   phosphate   synthase  from   Thermus   thermophilus   HB8:   implications   for   thermal   stability.   Acta  Crystallographica  Section  D-­Biological  Crystallography  2011,  67,  1054-­‐1064.    13.   Merz,   A.;   Knochel,   T.;   Jansonius,   J.   N.;   Kirschner,   K.,   The   hyperthermostable  indoleglycerol   phosphate   synthase   from   Thermotoga   maritima   is   destabilized   by  mutational   disruption   of   two   solvent-­‐exposed   salt   bridges.   Journal   of   Molecular  Biology  1999,  288  (4),  753-­‐763.    14.   Eberhard,   M.;   Tsai-­‐Pflugfelder,   M.;   Bolewska,   K.;   Hommel,   U.;   Kirschner,   K.,  Indoleglycerol   phosphate   synthase-­‐phosphoribosyl   anthranilate   isomerase:  comparison   of   the   bifunctional   enzyme   from   Escherichia   coli   with   engineered  monofunctional  domains.  Biochemistry  1995,  34  (16),  5419-­‐28.    

 

 

58  

15.   Kirschner,   K.;   Szadkowski,   H.;   Jardetzky,   T.   S.;   Hager,   V.,  Phosphoribosylanthranilate   Isomerase-­‐Indoleglycerol-­‐Phosphate   Synthase   from  Escherichia-­‐Coli.  Methods  in  Enzymology  1987,  142,  386-­‐397.    16.   Schneider,   B.;   Knochel,   T.;   Darimont,   B.;   Hennig,   M.;   Dietrich,   S.;   Babinger,   K.;  Kirschner,   K.;   Sterner,   R.,   Role   of   the   N-­‐terminal   extension   of   the   (betaalpha)8-­‐barrel   enzyme   indole-­‐3-­‐glycerol   phosphate   synthase   for   its   fold,   stability,   and  catalytic  activity.  Biochemistry  2005,  44  (50),  16405-­‐12.    17.  Hommel,  U.;  Eberhard,  M.;  Kirschner,  K.,  Phosphoribosyl  anthranilate  isomerase  catalyzes  a  reversible  amadori  reaction.  Biochemistry  1995,  34  (16),  5429-­‐39.    18.  Czekster,  C.  M.;  Neto,  B.  A.  D.;  Lapis,  A.  A.  M.;  Dupont,  J.;  Santos,  D.  S.;  Basso,  L.  A.,  Steady-­‐state  kinetics  of   indole-­‐3-­‐glycerol  phosphate  synthase  from  Mycobacterium  tuberculosis.  Archives  of  Biochemistry  and  Biophysics  2009,  486  (1),  19-­‐26.    19.  Schlee,  S.;  Dietrich,  S.;  Kurcon,  T.;  Delaney,  P.;  Goodey,  N.  M.;  Sterner,  R.,  Kinetic  Mechanism   of   Indole-­‐3-­‐glycerol   Phosphate   Synthase.   Biochemistry   2012,   52   (1),  132-­‐142.    20.  Wilmanns,  M.;   Priestle,   J.   P.;   Niermann,   T.;   Jansonius,   J.   N.,   Three-­‐dimensional  structure   of   the   bifunctional   enzyme   phosphoribosylanthranilate   isomerase:  indoleglycerolphosphate  synthase  from  Escherichia  coli  refined  at  2.0  A  resolution.  J  Mol  Biol  1992,  223  (2),  477-­‐507.    21.  Rothlisberger,  D.;  Khersonsky,  O.;  Wollacott,  A.  M.;  Jiang,  L.;  DeChancie,  J.;  Betker,  J.;  Gallaher,  J.  L.;  Althoff,  E.  A.;  Zanghellini,  A.;  Dym,  O.;  Albeck,  S.;  Houk,  K.  N.;  Tawfik,  D.  S.;  Baker,  D.,  Kemp  elimination  catalysts  by  computational  enzyme  design.  Nature  2008,  453  (7192),  190-­‐U4.      22.  Jiang,  L.;  Althoff,  E.  A.;  Clemente,  F.  R.;  Doyle,  L.;  Rothlisberger,  D.;  Zanghellini,  A.;  Gallaher,  J.  L.;  Betker,  J.  L.;  Tanaka,  F.;  Barbas,  C.  F.;  Hilvert,  D.;  Houk,  K.  N.;  Stoddard,  B.  L.;  Baker,  D.,  De  novo  computational  design  of  retro-­‐aldol  enzymes.  Science  2008,  319  (5868),  1387-­‐1391.    

 

 

 

59  

Chapter 3

Functional Identification of the General Acid and Base in the

Dehydration Step of IGPS Catalysis

[This Chapter was adapted from the paper entitled “Functional identification

of the general acid and base in the dehydration step of indole-3-glycerol

phosphate synthase catalysis” by Margot J. Zaccardi, Eric M. Yezdimer, and

David D. Boehr in The Journal of Biological Chemistry 2013.]

3.1 Abstract

The chemical mechanism for IGPS was proposed by Parry to proceed through two

intermediates in a series of condensation, decarboxylation, and dehydration steps. X-ray

crystal structures have suggested that Lys110 behaves as a general acid both in the

condensation and dehydration steps, but did not reveal an efficient pathway for the

reprotonation of this critical residue. Amino acid substitutions and kinetic experiments

presented in this chapter suggest an alternative mechanism whereby Lys110 acts as a

general acid in the condensation step, but another invariant residue, Lys53, acts as the

general acid in the dehydration step. These studies also indicate that the conserved

residue Glu51 acts as the general base in the dehydration step. The revised mechanism

effectively divides the active site into discrete regions where the catalytic surfaces

containing Lys110 and Lys53/Glu51 catalyze the ring closure (i.e. condensation and

decarboxylation) and dehydration steps, respectively.

 

 

60  

3.2 Introduction

The indole ring is a prevalent structure in biological systems and is found in

molecules that are relevant for many different industries including pharmaceuticals,

agriculture, and materials.1 Therefore, the synthesis of indoles remains a widely studied

area of research.2 New improvements for indole syntheses can be found through a better

understanding of the IGPS enzyme. The IGPS catalyzed reaction was previously

proposed to occur through two intermediates (I1, I2) in a series of condensation,

decarboxylation and dehydration steps (Figure 3.1).3, 4 Initiation of the enzyme-catalyzed

reaction is thought to begin with the protonation of the ketone in CdRP through general

acid catalysis by Lys110 (numbering according to ssIGPS), an absolutely conserved and

essential amino acid residue,4-6 which allows for electrophilic attack from the benzyl ring

that can drive the reaction forward to create the I1 intermediate; tandem mass

spectrometry has identified fragment patterms that were attributed to this intermediate.7

Following decarboxylation and the formation of the I2 intermediate, dehydration is

facilitated by a general acid and base (predicted to be Lys110 and Glu159, respectively),

to form the final product, IGP. Functional and structural studies are mostly consistent

with the currently accepted mechanism, but several unresolved issues remain, incuding

the proposition that Lys110 acts as the general acid in both the ring closure and

dehydration steps of the reaction. Not only is it atypical for an enzyme to use the same

residue for two separate proton donation steps, it also requires that Lys110 be

reprotonated in the middle of the catalytic reaction. An alternate hypothesis is that

another residue, such as Lys53, acts as the general acid in one of these steps.

 

 

61  

Figure 3.1: Proposed mechanism for catalysis by IGPS suggests that the reaction proceeds in three steps: condensation, decarboxlation, and dehydration, with two intermediates. The proposed general acid and base are Lys110 and Glu159. Problems with this mechanism include the need to reprotonate Lys110 between the first and third steps and the direct removal of an alkyl hydrogen from the pyrrole ring.

 

 

62  

Crystal structures of ssIGPS in complex with a reduced CdRP (rCdRP) analog

(i.e. a C2’ hydroxyl in place of the C2’ ketone) imply that the subtrate binds with the

enzyme in an extended, unproductive conformation, where the C1 and C2’ sites are

separated by about 4.5 Å, a distance too large for C-C bond formation.4 This finding

implies that CdRP must go through significant internal reorientation within the active site

after its activation by a general acid in order for condensation to occur. Interestingly, the

crystal structure of the ssIGPS:IGP binary complex shows that the anthranilate moiety is

docked into an adjacent hydrophobic pocket compared to crystal structures of the

IGPS:rCdRP complex, while the triosephosphate group remains secured in the same

position throughout the reaction. This result leads to the possibility that reorientation of

the substrate or intermediate may occur during catalysis. However, how or why this

reorientation occurs has not been resolved. A better understanding of this process would

be of utility in the industrial, biosynthetic production of new indole, the engineering of

new reactivity onto the IGPS active site, and the design of the most effective small

molecule inhibitors of IGPS for new antimicrobials.

One interaction that may be important in these conformational rearrangements of

the substrate is the salt bridge between Lys53 and the carboxylate moiety of the

anthranilate ring of CdRP; the enzyme is not able to catalyze the reaction with a substrate

analog lacking this carboxyl group.8 Lys53 is located on the highly dynamic β1α1 loop,

and Bruice and co-workers have proposed that the increased flexibility of this loop at

higher temperatures allows the substrate to form a more productive NAC for I1

formation.9, 10 The interaction between Lys53 and the carboxylate group may also be

involved in activating the C1 carbon for nucleophilic attack. The importance of Lys53

 

 

63  

was further demonstrated in activity assays of ssIGPS variants with substitutions at

Lys53 in ecIGPS, which demonstrated that Lys53 (Lys55 in ecIGPS) is essential to IGPS

catalysis.11

In this study, a series of kinetic experiments were performed in order to elucidate

the role of the highly conserved, charged residue in the active site of IGPS and allow for

a more complete understanding of the chemical mechanism for this enzyme. The

experiments reveal that Lys53 and Glu51 act as the general acid-base pair in the reaction,

catalyzing the dehydration of I2 to IGP product. This assignment differs from the

previously proposed assignments of either Glu159 or Glu210 as determined from

crystallography4 and MD simulations.9, 10 In light of this new assignment, we also

concluded that there are separate catalytic surfaces in IGPS that catalyze the ring closure

and dehydration; the I2 intermediate must undergo a reorientation during the reaction in

order for dehydration to occur.

3.3 Experimental Methods

3.3.1 Overexpression, Purification, and Kinetic Analysis of WT and Amino Acid

Substituted IGPS

Site-directed mutagenesis was performed to obtain Glu51Gln, Lys53Arg,

Lys53Gln, Glu159Gln, Arg182Ala, and Glu210Gln variants of ssIPGPS as described in

Chapter 2. All sequences for WT and mutant ssIGPS were confirmed through DNA

sequencing (Nucleic Acid Facility, Pennsylvania State University). The overexpression,

purification, and steady-state assays followed previously published protocols and were

performed as described in Chapter 2.4, 6, 12-14 The original WT ssIGPS overexpressed

 

 

64  

from the pET21b14 is N-terminal hexa-histidine tagged, but ssIPGS from pET26 is not.

Important to these studies, the His-tagged and non His-tagged versions of ssIGPS gave

essentially identical kinetic parameters (kcat(WT-hexahis)/kcat(WT-notag) = 1.2 ± 0.3).

All kinetic experiments were performed at 75 °C (unless otherwise noted). SVE,

SDKIE, and pH effects were determined by varying the buffer conditions of the original

assay as described in Chapter 2. Assays were performed at saturating substrate

concentrations of CdRP (800 nM for WT, 4000 nM for Glu210Gln, 8000 nM for

Glu51Gln, Lys53Arg, and Arg182Ala).

3.3.2 Overexpression and Purification of ε-13CH2-Lys Labeled ssIGPS

In order to selectively, isotopically label ssIGPS with ε-13CH2-Lysine, the enzyme

was overexpressed similarly to previously described.15 The following was dissolved into

950 mL water: 0.50 g alanine, 0.40 g aspartate, 0.40 g arginine, 0.05 g cystine, 0.40 g

glutamine, 0.65g glutamate, 0.55 g glycine, 0.10 g histidine, 0.23 g isoleucine, 0.23 g

leucine, 0.20 g 13C-ε-CH2-lysine (Cambridge Isotopes), 0.25 g methionine, 0.13 g

phenylalanine, 0.10 g proline, 0.50 g serine, 0.23 g threonine, 0.17 g tyrosine, 0.23 g

valine, 0.50 g adenine, 0.20 g thymine, 0.20 g cytosine, 0.50 g uracil, 1.50 g sodium

acetate, 1.50 g succinate, 0.50 ammonium chloride, 0.85 g sodium hydroxide (NaOH),

and 10.5 g dibasic potassium phosphate. This mixture was autoclaved and then allowed

to cool to room temperature. Then, 50 mL 40 % (w/v) glucose, 4 mL 1 M magnesium

sulfate, 1 mL 0.01 M iron chloride, and 10 mL of a vitamin mixture containing 2 mg

calcium chloride, 2 mg zinc sulfate, 2 mg manganese sulfate, 50 mg tryptophan, 50 mg

thiamine, 50 mg niacin, and 1 mg biotin was added. The pH was adjusted to

 

 

65  

approximately 7.3 and the solution sterile filtered.

Plasmid (pET21b) containing the his-tagged ssIGPS gene was transformed into

BL21-CodonPlus(DE3)-RIPL E. coli cells as described in Chapter 2. Fresh

transformations were used to inoculate 5 mL LB starter cultures, which were grown

overnight at 37 °C. 1 mL of this starter culture was used to inoculate 50 mL of the

selectively labeled media which was grown for approximately 20 hours at 37 °C. This

culture was used to inoculate the remainder of the media, which was grown at 37 °C until

the optical density (A600) was between 0.500 and 0.600 at which time the culture was

induced with IPTG and grown for 16 to 20 hours at 25 °C. All cell cultures contained 100

µg/mL Ampicillin. Cells were harvested by centrifugation (10,000 xg, 4 °C, 20 minutes)

and IGPS was purified and concentrated as previously described for His-tagged IGPS.

3.3.3 Preparation of rCdRP

CdRP was reduced to form the unreactive substrate analog, rCdRP through

selective reduction of CdRP with sodium borohydride (NaBH4). A solution of 5 M

NaBH4 was prepared in 0.05 M NaOH and added to a solution of CdRP such that the

ratio of NaBH4:CdRP was 1:3. The pH of the solution was adjusted with 0.05 M NaOH

until basic (pH 8 to 9). The solution was allowed to react for approximately one hour at

room temperature in the dark. Then, the pH was adjusted to approximately 5.0 with 1 N

hydrochloric acid.

rCdRP was purified by high performance liquid chromatography (HPLC) using a

Hypersil Gold PFP preparatory column (Thermo Scientific). The rCdRP was diluted by

approximately 50% with water and 50 µL was injected onto the column. The molecule

 

 

66  

was eluted using a gradient from 99% water, 1% tetrafluoroacetic acid (TFA) to 99%

acetonitrile, 1% TFA over 30 minutes with a flow rate of 2 mL/min. The fractions

containing purified rCdRP were identified using its absorbance at 327 nm and full

conversion to the reduced form was tested by adding a small portion to tmIGPS to

confirm that no turnover to IGP occured. The fractions were lyophilized to a powder and

reconstituted in D2O to a concentration of 5 mM.

3.3.4 13C-HSQC Experiments on ssIGPS

After purification and concentration of the sample, IGPS was diluted to

approximately 1 mM and exchanged into buffer containing 25 mM potassium phosphate

pH 7.0 (pD 6.6), 75 mM potassium chloride, 1 mM EDTA, 1 mM dithiothreitol (DTT),

0.02% sodium azide (prepared in 100% D2O) using ZEBA desalting spin columns

(Thermo Scientific). Standard 1H-13C-HSQC (heteronuclear single quantum coherence)

experiments were obtained at 306 K on a Bruker AV-III-500 MHz spectrometer. pH was

adjusted for the titration by adding small volumes (< 5µL) of 0.5 M NaOH prepared in

D2O. The pKas were determined by plotting the change in chemical shift of hydrogen for

each peak and fitting to the equation:

€

δ = C /(1+10pKa − pH ) (3.1)

where δ is 1H chemical shift and C is the pH-independent rate value.

 

 

67  

3.4 Results

3.4.1 Determination of the Rate-Determining Step of ssIGPS Catalysis

There are five highly conserved, charged residues in the active site of IGPS:

Glu51, Lys53, Lys110, Glu159, Arg182, and Glu210. Previous kinetic experiments in

ecIGPS indicate that these residues are all important for catalysis, but that study was not

able to delineate the particular role for each residue. Using similar methodology as in

Chapter 2 (Figure 2.2), SVE and SDKIE experiments were used to define the specific

role for these residues. SVE report on steps that are diffusion-controlled (substrate

binding, product release, and large conformational changes), whereas SDKIEs report on

chemical steps involving proton transfer. At 75 °C, the rate-determining step of WT

ssIGPS is the ring closure, as evidenced by the presence of an SDKIE and lack of an

SVE, whereas at 25 °C, product release is rate-determining as evidenced by an SVE of

approximately one. The same methodology was used to examine the rate-determining

step for the ssIGPS variants.

3.4.2 Analysis of Lys53 Indicates its Role as a General Acid

The assignment of Lys110 as the general acid in both steps of the reaction was based on

crystal structures of the enzyme, but there is no obvious mode of reprotonation, leading to

our hypothesis that an alternate residue is responsible for donating a proton in one of

these steps. Excluding Lys110, the only remaining cationic residues in the active site

capable of performing the role of general acid are Lys53 and Arg182. Therefore, enzyme

variants with amino aids substitutions at these residues were kinetically examined (Table

3.1). The Arg182Ala substitution had only a minor effect on the catalytic turnover, kcat,

 

 

68  

Table 3.1: Steady-state kinetics (at 75 °C) demonstrates that the dehydration step of IGPS catalysis occurs through the general acid and base Lys53 and Glu51, respectively.

aDue to the low activity of Lys53Gln, a full steady-state curve was not possible. The kcat reported is the kobserved at saturating substrate concentrations bActivity of Lys110Arg and Glu159Gln variants was below the limit of detection. The value reported is the lowest kobs able to be measured through the fluorescence assay

IGPS

Variant

akcat (s-1) aKM (nM) kcat/KM

(x106 M-1s-1)

SDKIE

(kH2O/kD2O)

SVE

WT 0.67 ± 0.03 44 ± 9 15 3.6 ± 0.3 -0.2 ± 0.1

Glu51Gln 0.007 ± 0.001 13.0 ±4.4 0.54 0.9 ± 0.1 0.16 ± 0.2

Lys53Arg 0.06 ± 0.01 2440 ± 3 0.03 1.0 ± 0.1 0.2 ± 0.1

Lys53Gln 0.001 ± 0.001 n.d.a n.d.a n.d.a n.d.a

Arg182Ala 0.41 ± 0.07 1062 ± 499 0.38 1.1 ± 0.6 0.7 ± 0.3

Glu210Gln 0.97 ± 0.07 386 ± 96 2.5 1.0 ± 0.3 0.7 ± 0.2

Lys110Arg < 1.5 x 10-5 b n.d.b n.d.b n.d.b n.d.b

Glu159Gln < 1.5 x 10-5 b n.d.b n.d.b n.d.b n.d.b

 

 

69  

which suggests that this residue is not directly involved in the chemical steps of the

reaction. Conversely, this variant showed a substantial increase in KM (~20 fold). This

result, along with the considerable SVE and loss of SDKIE for this variant, implicates its

importance in substrate binding.

To test the importance of the positive charge on Lys53 and its ability to act as a

general acid, we assayed Lys53Arg and Lys53Gln ssIGPS variants; Arg retains the

positive charge and some ability to donate a proton, whereas Gln does not have either

function but would still be able to participate in CdRP binding through a hydrogen bond

interaction with the carboxylate moiety. Since Lys53 is known to participate in CdRP

binding, changes in substrate binding were expected for these variants since substitutions

at this position would create less optimal interactions with CdRP. The Lys53Arg

substitution led to a decrease in CdRP affinity, with a 55-fold increase in KM (Table 3.1).

In addition, there was a 10-fold decrease in catalytic turnover (kcat) for Lys53Arg ssIGPS,

which is consistent with a direct role for Lys53 in the chemical steps of IGPS catalysis.

The Lys53Gln substitution had an even more deleterious effect on IGPS catalysis,

resulting in an over 500 fold-decrease in activity, even at the very high CdRP

concentration of 12 µM (50 times KM for WT IGPS). This finding indicates that the

positive charge and likely the ability of Lys53 to act as a proton donor are critical for

IGPS catalysis.

SVE, SDKIE, and pH effects further established the role of Lys53 in the

chemistry of the IGPS catalyzed reaction. Lys53Arg ssIGPS had a slightly larger SVE

than WT ssIGPS, which is consistent with a decrease in the rate of substrate binding or

product release, causing these processes to make a slight contribution to the overall rate.

 

 

70  

The pH and SDKIE results indicate the specific role for Lys53 in reaction chemistry. The

bell-shaped pH curve for WT IGPS is consistent with acid-base catalysis (Figure 3.2),

but Lys53Arg displays a substantially higher pKa for the descending limb compared to

the WT enzyme (Table 3.2), closer to what is expected for an Arg residue. This result,

together with the finding that the Gln substitution is much more detrimental than Arg

(Table 3.1), suggest that Lys53 is behaving as a general acid during the reaction. The loss

of an SDKIE in Lys53Arg IGPS compared to the WT enzyme indicates that the rate-

determining step of the Lys53Arg ssIGPS catalyzed reaction is the dehydration rather

than the ring closure; the ring closure is expected to be isotope sensitive due to the proton

donation from Lys110, but the dehydration step is proposed to be isotope insensitive (see

Appendix for kinetic analysis). This proposal is also consistent with the decreased

SDKIE in WT ssIGPS at higher pH values, and the finding that the SDKIE arises from a

single proton transfer event.6

 

 

71  

Figure 3.2: The pH rate profiles suggest that Lys53 and Glu51 act as the general acid and base, respectively in the dehydration step of IGPS catalysis. Shown are the pH curves for IGPS (a) WT (b) Lys53Arg and (c) Glu51Gln at 25 °C (left) and 75 °C (right).

 

 

72  

Table 3.2: pKa values for WT ssIGPS and Lys53Arg and Glu51Gln variants identify Lys53 and Glu51 as the general acid and base in ssIGPS catalysis. Enzyme Variant Temperature (°C) pKa1 pKa2

WT 25 5.4 ± 0.2 8.9 ± 0.2

75 5.6 ± 0.2 8.7 ± 0.1

Lys53Arg 25 6.9 ± 0.1 > 9.5

75 7.1 ± 0.1 > 9.5

Glu51Gln 25 5.7 ± 0.2 n/a

75 6.51 ± 0.3 n/a

 

 

73  

3.4.3 13C-Lys NMR to Determine pKas for Lysine Residues in IGPS

In order to conclusively determine which residues in the active site were

responsible for the pKa2 observed in the steady-state pH experiments, 13C-1H HSQC

nuclear magnetic resonance (NMR) experiments were performed on an IGPS sample

labeled with 13C-Lys. Eighteen distinct resonances were identified in the spectrum

(Figure 3.3), which is consistent with the seventeen lysine residues in ssIGPS, though

some of these resonances may represent different protons from the same lysine, such as

peaks 2 and 3 which show the same 13C chemical shift but different 1H chemical shifts.

A pH titration was performed on the ssIGPS enzyme in order to assign pKa values

to each resonance in the spectrum and correlate the pKa in the steady-state assay to a

particular residue in the enzyme. The pKa values were measured by tracking the change

in chemical shift (Figure 3.4) and plotting chemical shift vs pH for each peak (Figure

3.5) determined using NMR were significantly higher than the pKa of the descending

limb for the pH profile (Table 3.3). The pH rate dependence experiments indicate that at

high pH values (>9.5) the enzyme begins to denature. It is likely that the pH titration and

pKa values are not valid due to this denaturation. In the pH range where ssIGPS is known

to be stable there was no change in chemical shift. Therefore, it is more likely that the

change in chemical shift at higher pH is due to denaturation, rather than the deprotonation

of the lysine residues.

To test whether the introduction of D2O in the NMR experiments was affecting

the titration, the pH dependence of the enzyme activity was performed in assay buffer

prepared in D2O. The results (Figure 3.5) showed that there was no difference in the pKa

values for the kinetic pH dependence in D2O versus H2O.

 

 

74  

Figure 3.3: 1H-13C HSQC on ε-13CH2-Lys labeled ssIGPS shows seventeen resonances which is consistent with the number of lysine residues in the enzyme

 

 

75  

Figure 3.4: Overlay of 1H-13C-HSQC Spectra for ssIGPS at pH 7.0 (black) and pH 10.5 (red). At high pH, the changes in the spectrum are likely caused by denaturation of ssIGPS.

 

 

76  

 

Figure 3.5: A representative plot of chemical shift versus pH for peak #2. The pKa value associated with this curve is 11.45 ± 0.14, although the change in chemical shift likely reflects denaturation of the enzyme rather than deprotonation of the lysine.

 

 

77  

Table 3.3: pKa values determined by 1H-13C HSQC on ε-13CH2-Lys labeled ssIGPS are not in agreement with the pKa for the pH dependence of the enzymatic reaction determined by steady-state kinetics.

Resonance   pKa     Resonance   pKa    

0   10.98  ±  0.34   9   10.94  ±  1.43  

1   11.27  ±  0.16   10   9.63  ±  0.16  

2   11.45  ±  0.14   11   10.86  ±  0.18  

3   11.42  ±  0.13   12   11.43  ±  0.18  

4   11.05  ±  0.10   13   11.88  ±  0.95  

5   11.38  ±  0.12   14   11.34  ±  0.14  

6   11.37  ±  0.13   15   11.06  ±  0.14  

7   11.36  ±  0.12   16   11.53  ±  0.20  

8   11.30  ±  1.05   17   10.72  ±  0.14  

 

 

78  

Figure 3.5: pH dependence of the ssIGPS catalyzed reaction performed in H2O (pKa1 5.7 ± 0.1, pKa2 8.7 ± 0.1), shown in blue, and D2O (pKa1 5.3 ± 0.2, pKa2 8.9 ± 0.2), shown in green, display little difference in pKa values. Therefore, the use of D2O in NMR experiments does not explain the discrepancy in pKa values between the two methods.

 

 

79  

In addition to the NMR titration results not aligning with the steady-state assays,

assignment of the resonances in the spectrum also proved to be problematic. Assignments

were attempted by performing NMR experiments with amino acid substituted variants of

ssIGPS; it was expected that deletion of a Lys would lead to the loss of a resonance in the

spectrum. The Lys53Arg sample was not stable in solution at high concentrations (~1

mM) and much of the sample precipitated during these experiments. The spectrum for

Lys53Arg ssIGPS resembles that of WT ssIGPS, but the low resolution made it

impossible to assign the residue. While it is still unclear which resonance belongs to

Lys53Arg (Figure 3.6), it is clear that the Lys53Arg substitution changes the chemical

environment of several of the resonances compared with WT enzyme at the same pH.

In a final attempt to assign the residues, 1H-13C-HSQC experiments were

performed in the presence of rCdRP. The addition of this substrate analog may change

the chemical environment of some residues in the enzyme, leading to a more dispersed

spectrum. Unfortunately, even with high concentrations of rCdRP (5 times [IGPS]) the

spectrum did not show any significant differences from the ligand free WT ssIGPS

spectrum. Due to the poor alignment of the NMR data to the steady-state data, along with

the inability to assign resonances in the NMR spectrum, these experiments were

unsuccessful and did not provide any results that are useful in the interpretation of the

kinetic studies.

 

 

 

80  

Figure 3.6: Overlay of 1H-13C HSQCs of Lys53Arg (red) and WT ssIGPS (black) labeled with ε-CH2-Lys. Due to the low resolution of Lys53Arg ssIGPS spectrum, and its the poor alignment to WT, Lys53 and other resonances remain unassigned.

 

 

81  

3.4.4 Analysis of Glu51 Identifies its Role as the General Base

Both Glu159 and Glu210 were previously proposed to act as the general base in

the dehydration of I2 to IGP. Glu159 was proposed because it is essential to the reaction,

the analogous amino acid substitution in ecIGPS, Glu163Gln, showed a 540-fold

decrease in kcat,5 and because Glu159 would appear to be appropriately positioned if

Lys110 is the general acid in the dehydration. Glu210 was alternately predicted as the

general base through MD simulations, as both Gu159 and Glu210 are both a similar

distance from the C1’ of the substrate where attack by the general base is proposed to

occur.9, 10 Since our results demonstrate that Lys53, rather than Lys110, is acting as the

general acid in the dehydration, it is less likely that Glu159 or Glu210 could be

performing the role of general base due to their distal location from Lys53, which would

prevent appropriate contact of I2 with both Lys53 and Glu159/Glu210.4, 12, 16

To probe the role of these residues, Glu159Gln and Glu210Gln variants were

examined. Similar to the analogous change in ecIGPS, the Glu159Gln substitution in

ssIGPS led to a substantial decrease in catalytic activity (kobs< 1.5 x 10-5 s-1). This change

in activity is on the same order of magnitude that was previously seen for the Lys110Arg

variant,6 which likely indicates that Glu159 and Lys110 are operating on the same

chemical step of the reaction (ring closure), which is also consistent with previous

proposals by Bruice and coworkers.10 The Glu210Gln variant did not show any decrease

in kcat, indicating that Glu210 is not involved directly in the chemistry of the reaction.

However, the loss of binding affinity (four-fold increase in KM) for the Glu210Gln

variant suggests that Glu210 plays a role in substrate binding.

Besides Glu159 and Glu210, Glu51 is the only other conserved residue in the

 

 

82  

active site that may be performing this function.4 Kinetic analysis of the Glu51Gln

variant shows a 100-fold decrease in activity compared to WT enzyme (Table 3.1). The

kinetic parameters are on the same order of magnitude as those seen for Lys53Arg, which

is expected for variants that affect the same step of the kinetic mechanism. The Glu51Gln

variant also shows a complete loss of the SDKIE for kcat, as was also observed for the

Lys53Arg variant, which indicates that this residue is also involved in the dehydration

step of the reaction. Most telling was the pH dependence of Glu51Gln activity; the

variant displays a complete loss in the ascending limb, which is indicative of a loss in

general base activity for this variant (Figure 3.2). Altogether our results implicate

Lys53/Glu51 as the general acid/base pair responsible for catalyzing the dehydration step

of the reaction.

3.5 Discussion

In this study, we have delineated the roles for the highly conserved, charged

residues present in the active site of ssIGPS (Figure 3.7). Consistent with previous

proposals, Lys110 acts as the general acid in the ring closure step, and is likely aided by

Glu159. Our studies indicate Lys53 and Glu51 are responsible for catalyzing the

dehydration step as the general acid and base, respectively. Additionally, Arg182 and

Glu210 participate in substrate binding.

The newly identified roles for Glu51 and Lys53 are at odds with the crystal

structure of the IGPS:IGP complex; in the crystallized conformation, neither Lys53 nor

Glu51 is appropriately positioned to facilitate catalysis. However, a rotation through the

C3’-C4’ bond of the ribose chain relocates the amine group of I2 and the C2’ hydroxyl

 

 

83  

Figure 3.7: The assigned role for the conserved and charged residues in the active site of IGPS. The ring closure is catalyzed by the general acid, Lys110, and Glu159 (blue). The dehydration is catalyzed by the general acid, Lys53, and general base, Glu51 (yellow). Arg182 and Glu210 (orange) are involved in susbtrate binding.

 

 

84  

within contact distance of Glu51 and Lys53, respectively. This rotation is reasonable if

the previously suggested internal bond rotations of CdRP required to close the initial 4.5

Å distance between the C1 and C2’ for the ring closure, the two previously identified

binding pockets for the anthranilate moiety of the substrate, and the weak

crystallographic electron density of the ribulose moiety in the bound protein are

considered. Additionally, there likely is sufficient thermal flexibility of the substrate

within the active site to reorient the I2 intermediate into a appropriate position with

respect to both the Glu51 and Lys53 residues.

Based on this study, we propose a revised mechanism for IGPS catalysis (Figure

3.8). Following the ring closure, internal rotations in the ribose ring reposition the I2

intermediate in a position where Lys53 is in close proximity to the C2’ hydroxyl, and

Glu51 is in close proximity to the amine hydrogen (Figure 3.9). During the final step, the

lone pair of electrons on Glu51 attacks the hydrogen bond of the amine group. The

original mechanism proposed the general base instead attacked the C1’ hydrogen;

however, the loss of a hydrogen from an amine (pKa ~21) is much more reasonable that

from a carbon (pKa ~44 to 51). This action starts an electron cascade that ends with the

proton donation by Lys53 to the hydroxyl of C2’, releasing H2O and forming the product,

IGP. The reaction leaves the side chain of Lys53 neutralized, weakening the electrostatic

attraction between the protein and the product, which likely acts as an important trigger

event for product release.

 

 

85  

Figure 3.8: The modified mechanism of ssIGPS catalysis utilizes Lys53 and Glu51 as the general acid and base pair in the dehydration step of the reaction. Additionally, the general base now attacks the amine hydrogen rather than the previously suggested alkyl hydrogen.

 

 

86  

Figure 3.9: Rotation about the C3’-C4’ bond of ribose chain is required for the dehydration step in IGPS catalysis. Crystal structure of ssIGPS:IGP (Top) (PDB: 1A53) complex shows the ligand bound in the active site such that Lys53 and Glu51 are not properly positioned for catalysis. The ribose chain must rotate (Bottom) to reposition the intermediate in the second binding pocket and allow for dehydration to form IGP.

 

 

87  

Mapping this revised mechanism on to the active site reveals two distinct regions

of function (Figure 3.10). The first region contains the surfaces formed by the

combination of Lys110 and Glu159 and it is this region that is likely responsible for the

ring closure and decarboxylation step of the catalytic pathway. The second region is

located adjacent to the first and includes Glu51 and Lys53. This region is responsible for

completing the dehydration in the second step of the mechanism. After the dehydration

reaction, Lys53 is rendered neutral likely causing a decrease in binding affinity that leads

to the release of IGP from the enzyme. Additionally, the binding site for the dehydration

step is located closer to where the product will exit the active site following the reaction,

and it is likely that this aids in the release of IGP as well.

 

 

88  

Figure 3.10: Surface rendering of the IGPS binding pocket shows two distinct active sites for catalysis. In the first site (blue), Lys110 and Glu159 catalyze the ring closure step deep within the pocket. The intermediate then transitions to the second site (yellow), which is closer to where product exits the binding pocket, where Lys53 and Glu51 catalyze the dehydration step. (PDB: 1A53).

 

 

89  

3.6 Conclusions

The production of indole, tryptophan, and their derivatives is both a critical

biological process and one of extreme utility in the chemical and pharmaceutical

industries. The new experimental data on ssIGPS WT and variants presented in this

Chapter supports a modified catalytic mechanism for IGPS, whose previously proposed

mechanism had been standing for the past forty years. This mechanism divides the active

site into discrete regions where the first catalytic surface, containing Lys110 and Glu159,

catalyzes the ring closure step, and the second, containing and Lys53 and Glu51,

catalyzes the dehydration step. Since IGPS is a validated antimicrobial target, this new

mechanism can be used in the rational drug design of original IGPS inhibitors that target

one or both of the catalytic sites. IGPS has also been used as a superior starting scaffold

in the development of novel enzymes. Engineered IGPS enzymes should find utility in

the development of new synthetic schemes for novel indole derivatives that may find

medical and agricultural use.

3.7 References

1.  Barden,  T.  C.,  Indoles:  Industrial,  agricultural  and  over-­‐the-­‐counter  uses.  Topics  in  Hetercyclic  Chemistry  2011,  26,  31-­‐46.    2.  Humphrey,  G.  R.;  Kuethe,  J.  T.,  Practical  methodologies  for  the  synthesis  of  indoles.  Chemical  Reviews  2006,  106  (7),  2875-­‐2911.    3.  Smith,  M.;  March,   J.,  March's  advanced  organic  chemistry   :  reactions,  mechanisms,  and  structure.  5th  ed.;  John  Wiley:  New  York,  2001;  p  xviii,  2083.    4.  Hennig,  M.;  Darimont,  B.  D.;  Jansonius,  J.  N.;  Kirschner,  K.,  The  catalytic  mechanism  of   indole-­‐3-­‐glycerol   phosphate   synthase:   crystal   structures   of   complexes   of   the  enzyme   from   Sulfolobus   solfataricus   with   substrate   analogue,   substrate,   and  product.  J  Mol  Biol  2002,  319  (3),  757-­‐66.  

 

 

90  

5.  Darimont,  B.;  Stehlin,  C.;  Szadkowski,  H.;  Kirschner,  K.,  Mutational  analysis  of  the  active   site   of   indoleglycerol   phosphate   synthase   from   Escherichia   coli.   Protein  Science  1998,  7  (5),  1221-­‐1232.    6.   Zaccardi,   M.   J.;   Mannweiler,   O.;   Boehr,   D.   D.,   Differences   in   the   catalytic  mechanisms  of  mesophilic   and   thermophilic   indole-­‐3-­‐glycerol   phosphate   synthase  enzymes   at   their   adaptive   temperatures.   Biochemical   and   Biophysical   Research  Communications  2012,  418  (2),  324-­‐329.    7.  Czekster,  C.  M.;  Lapis,  A.  A.  M.;  Souza,  G.  H.  M.  F.;  Eberlin,  M.  N.;  Basso,  L.  A.;  Santos,  D.   S.;   Dupont,   J.;   Neto,   B.   A.   D.,   The   catalytic   mechanism   of   indole-­‐3-­‐glycerol  phosphate   synthase   (IGPS)   investigated  by   electrospray   ionization   (tandem)  mass  spectrometry.  Tetrahedron  Letters  2008,  49  (41),  5914-­‐5917.    8.   Smith,   O.   H.;   Yanofsky,   C.,   1-­‐(ortho-­‐carboxyphenylamino)-­‐1-­‐deoxyribulose   5-­‐phosphate,   a   new   intermediate   in   the   biosynthesis   of   tryptophan.   Journal   of  Biological  Chemistry  1960,  235  (7),  2051-­‐2057.    9.   Mazumder-­‐Shivakumar,   D.;   Kahn,   K.;   Bruice,   T.   C.,   Computational   study   of   the  ground   state   of   thermophilic   indole   glycerol   phosphate   synthase:   Structural  alterations   at   the   active   site   with   temperature.   Journal   of   the   American   Chemical  Society  2004,  126  (19),  5936-­‐5937.    10.  Mazumder-­‐Shivakumar,  D.;  Bruice,  T.  C.,  Molecular  dynamics  studies  of  ground  state   and   intermediate   of   the   hyperthermophilic   indole-­‐3-­‐glycerol   phosphate  synthase.   Proceedings   of   the   National   Academy   of   Sciences   of   the   United   States   of  America  2004,  101  (40),  14379-­‐14384.    11.  Eberhard,  M.;  Kirschner,  K.,  Modification  of  a  catalytically   important  residue  of  indole-­‐3-­‐glycerol  phosphate  synthase   from  Escherichia  coli  Febs  Letters  1989,  245  (1-­‐2),  219-­‐222.    12.   Hennig,   M.;   Darimont,   B.;   Sterner,   R.;   Kirschner,   K.;   Jansonius,   J.   N.,   2.0   A  structure   of   indole-­‐3-­‐glycerol   phosphate   synthase   from   the   hyperthermophile  Sulfolobus  solfataricus:  possible  determinants  of  protein  stability.  Structure  1995,  3  (12),  1295-­‐306.    13.  Merz,  A.;  Yee,  M.  C.;  Szadkowski,  H.;  Pappenberger,  G.;  Crameri,  A.;  Stemmer,  W.  P.  C.;   Yanofsky,  C.;  Kirschner,  K.,   Improving   the   catalytic   activity  of   a   thermophilic  enzyme  at  low  temperatures.  Biochemistry  2000,  39  (5),  880-­‐889.    14.   Schneider,   B.;   Knochel,   T.;   Darimont,   B.;   Hennig,   M.;   Dietrich,   S.;   Babinger,   K.;  Kirschner,   K.;   Sterner,   R.,   Role   of   the   N-­‐terminal   extension   of   the   (betaalpha)8-­‐barrel   enzyme   indole-­‐3-­‐glycerol   phosphate   synthase   for   its   fold,   stability,   and  catalytic  activity.  Biochemistry  2005,  44  (50),  16405-­‐12.  

 

 

91  

15.  Muchmore,  D.  C.;  McIntosh,  L.  P.;  Russell,  C.  B.;  Anderson,  D.  E.;  Dahlquist,  F.  W.,  Expression   and   N-­‐15   labeling   of   proteins   for   proton   and   N-­‐15   nuclear-­‐magnetic  resonance.  Methods  in  Enzymology  1989,  177,  44-­‐73.    16.  Wilmanns,  M.;   Priestle,   J.   P.;   Niermann,   T.;   Jansonius,   J.   N.,   Three-­‐dimensional  structure   of   the   bifunctional   enzyme   phosphoribosylanthranilate   isomerase:  indoleglycerolphosphate  synthase  from  Escherichia  coli  refined  at  2.0  A  resolution.  J  Mol  Biol  1992,  223  (2),  477-­‐507.  

 

 

92  

Chapter 4

The Role of Active Site Loops in Catalysis by IGPS

4.1 Abstract

Substrate binding, product release, and likely chemical catalysis in the tryptophan

biosynthetic enzyme indole-3-glycerol phosphate synthase (IGPS) are controlled through

the structural dynamics of the β1α1 loop. We have probed the role of the nearby β2α2

loop and the interaction between the β1α1 and β2α2 loops using amino acid substitutions

and kinetic analysis to gain insight into how loop-loop interactions regulate IGPS

catalysis. Our results indicate that interactions between co-evolving residues Arg54 and

An90 on these loops are important in coordinating the multistep reaction in IGPS, and

can modulate the general acid activity of the absolutely conserved Lys53 on the β1α1

loop.

4.2 Introduction

The IGPS structure consists of a (β/α)8-barrel, or TIM-barrel fold, the most

common fold found in structural biology. The fold typically consists of eight parallel β-

strands in the shape of a wheel surrounded by eight α-helices. TIM-barrel proteins are

catalytically diverse,1 but the structural elements are highly conserved, and most of the

chemically relevant amino acids residues are located on the inside of the β-barrel on the

C-terminal end of the β-strands or the loops connecting β-strands to α-helices (βα loops).

These βα loops are shown to be particularly important in the proper function of the

enzyme. In addition to containing many of the active site residues that participate in the

 

 

93  

reaction, interactions between loops have been implicated in the function of many (β/α)8-

barrel enzymes2-4 including TIM where the structural dynamics of the β6α6 loop allows

for substrate binding and product release.5 Bioinformatics approaches and various

computational methods, including MD simulations, have also implicated an important

role for several loops in the (β/α)8-barrel enzyme IGPS,6-8 though the functional role of

these loops has not, until now, been investigated experimentally.

SCA combined with MD simulations identified a potentially important interaction

between residues on the β1α1 and β2α2 loops in IGPS.8 SCA identifies residue positions

that co-vary, and proposes that these residues are important in maintaining structure,

allowing proper protein folding, or more directly facilitating proper enzyme function and

catalysis.9 When used in conjunction with MD simulations, SCA can identify

functionally important amino acid pairs that are not absolutely conserved through all

species but whose interaction is conserved and whose coordinated motions may be

important in enzyme catalysis. A similar methodology also identified other co-varying

amino acids on IGPS, and mutational studies confirmed that these residues are

catalytically and/or structurally important.10 The co-varying residues in the β1α1 and

β2α2 loops may be especially important considering the functional role of the β1α1 loop

in IGPS catalysis.

Specifically, SCA-MD analysis suggests that interactions between Arg54 and

Asn90 on the β1α1 and β2α2 loops are important for enzyme function (Figure 4.1).

Structural comparison of the substrate-bound and product-bound forms of IGPS, together

with the structural modeling of the intermediate in the active site shows two distinct and

adjacent hydrophobic pockets, suggesting that the anthranilate group of the substrate

 

 

94  

Figure 4.1: The catalytically important residues in the dehydration step of IGPS are found on the β1α1 and β2α2 loops, which interact through a hydrogen bond between Arg54 and Asn90. (a) The ssIGPS catalyzed reaction has two distinct binding pockets for the two reaction steps. In step one, Lys 110 (cyan) initiates the ring closure and decarboxylation. Following the formation of the intermediate, the anthranilate moiety is transferred to the second site where Lys53 and Glu51 (yellow) act as the active site acid and base. The role of the β1α1 and β2α2 loops (blue) including the interaction between Arg54 and Asn90 is examined herein. (b) This interaction is thought to have functional significance in IGPS since it is coevolving amongst IGPS species and exhibits correlated motion in MD simulations. This interaction is in close proximity to the conserved residues Lys53 and Phe89.

 

 

95  

moves from one hydrophobic pocket to another during catalysis.11, 12 It is important to

note that the conserved Phe89 on the β2α2 loop comprises part of this anthranilate

binding pocket through π-π interations with the CdRP substrate. There is also a second

coevolving interaction between residues on the β2α2 loop through Glu85 and Asn90. We

hypothesize that the interaction between Arg54 and Asn90 (and perhaps Glu85) may help

to coordinate conformational changes in both the loops and the substrate or intermediate

itself.

In this Chapter, the importance of the β2α2 loop and its interaction with the β1α1

loop in ssIGPS catalysis was examined through the analysis of Arg54, Asn90, Glu85, and

Phe89 variants. The results indicate that the interaction between the two active site loops

in ssIGPS is important for proper function of the general acid/base pair, Lys53/Glu51, in

the dehydration step. It may also be involved in coordinating the transition of the

intermediate between the two binding sites during catalysis. These results provide insight

into the coordination of the multiple step reaction catalyzed by IGPS.

4.3 Experimental Methods

Site-directed mutagenesis of ssIGPS variants (Arg54Ala, Phe89Ala, Asn90Ala,

Asn90Gln. Arg54Lys, Arg54Ala/Asn90Ala, and Arg54Lys/Asn90Gln) was performed

using the QuikChange Lightning® Site-Directed Mutagenesis Kit (Agilent Technologies)

and appropriate primers as described in previous Chapters. All sequences (WT and

mutant) were confirmed through DNA sequencing (Nucleic Acid Facility, Pennsylvania

State University). Overexpression, purification, and steady-state kinetic analyses of WT

and variant enzymes were performed as previously described (see Chapter 2). 13,14

 

 

96  

Thermal inactivation experiments for ssIGPS enzymes (non His-tagged, WT and

amino acid substituted) were performed by incubating the stock enzyme solution at 90 °C

for up to 30 minutes. Aliquots were removed every three minutes and the enzyme activity

was assayed in duplicate at 50 °C with a saturating CdRP concentration (800 nM). The

data was fit to a linear regression of relative rate (compared to the rate prior to

incubation) versus incubation time in seconds. The first-order rate constant of thermal

inactivation, kinact, is equal to the slope of this linear fit.

Circular dicroism (CD) experiments were performed out on a Jasco J-810

Spectropolarimeter from 250 nm to 190 nm with 1 nm intervals and a 1 nm bandwidth.

The experiments were performed in 10 mM potassium phosphate pH 7.0 with an enzyme

concentration of 1.7 µM.

4.4 Results

4.4.1 Investigation of Phe89 on the β2α2 Loop Identifies its Role in IGPS Chemistry

Since we are interested in the role of active site loops in IGPS catalysis, and

previous studies resolved the role of the conserved residues on the β1α1 loop and

adjecent β1-strand (Lys53 and Glu51), we examined the role of the essential and

invariant Phe89 residue located on the β2α2 loop with an alanine substitution. Phe89 is

on the β2α2 loop and makes a π-π interaction with the anthranilate moiety of CdRP.

Therefore, it is not surprising that the Phe89Ala amino acid substitution also led to an

approximately 24-fold increase in KM (Table 4.1). Interestingly, the substitution led to an

eleven-fold decrease in kcat associated with this variant, which suggests that this residue

 

 

97  

Table 4.1: Steady-state kinetic parameters of ssIGPS WT and loop variants indicate an important role for the β1α1 and β2α2 loop interaction in catalysis Variant Temp

(°C) kcat s-1 KM (nM) kcat/KM

(x 106 M-1s-1)

(kcat)WT/(kcat)mutant

SVE SDKIE

WT 25 0.16 ± 0.02 74 ± 38 2.2 1.0 ±0.2 1.2 ± 0.2

37 0.42 ± 0.04 88 ± 47 4.8 0.6 ±0.3 5.8 ± 0.1 75 0.67 ± 0.03 44 ± 9 15 -0.2 ±0.1 3.6 ± 0.3 Phe89Ala 75 0.06 ± 0.01 1200 ±

700 0.05 11 -0.23 ±

0.07 1.0 ± 0.2

Arg54Ala 25 0.16 ± 0.01 145 ± 36 2.2 1.0 1.3 ± 0.4 n.d. 37 0.35 ± 0.04 95 ± 41 3.7 1.2 0.5±0.1 5.0 ± 1.0

75 1.2 ± 0.1 74 ± 13 16 0.55 -0.2 ±0.1 1.2 ± 0.2 Asn90Ala 25 0.04 ± 0.03 36 ± 21 1.0 4.7 0.7 ± 0.3 1.0 ± 0.3 37 0.17 ± 0.02 74 ± 31 2.3 2.5 0.06

±0.1 5.2 ± 0.4

75 0.55 ± 0.02 79 ± 10 7 1.2 -0.2 ±0.1 1.6 ±0.2 Arg54Ala/Asn90Ala

37 0.19 ± 0.06 23 ±12 8.2 2.2 0.0 ±0.3 2.2 ± 0.3

75 0.51 ± 0.08 107 ± 49 4.7 1.3 -0.1 ±0.1 1.0 ±0.2 Glu85Ala 37 0.26 ± 0.03 140 ± 46 1.9 1.6 0.8 ± 0.2 2.2 ± 0.6 75 0.85 ± 0.07 371 ± 81 2.7 0.8 0.2 ± 0.2 1.6 ± 0.4 Arg54Lys 37 0.31 ± 0.02 84 ± 17 3.7 1.3 0.4 ± 0.3 2.9 ±0.5 75 0.66 ± 0.13 379 ±

200 9.3 1.0 0.01 ±

0.1 1.8 ± 0.3

Asn90Gln 37 0.30 ±0.02 132 ±31 2.3 1.4 0.4 ± 0.1 4.1 ± 0.8 75 0.89 ±0.06 117 ±26 7.6 0.75 0.3 ± 0.1 1.7 ± 0.2 Arg54Lys/Asn90Gln

37 0.44 ± 0.02 155 ± 29 2.8 0.95 0.2 ± 0.1

3.5 ± 0.8

75 1.7 ± 0.1 462 ± 89 3.6 0.40 -0.1 ± 0.1

0.97 ± 0.4

 

 

98  

may also be involved in the chemistry of the reaction.

To gain additional insight into the binding and chemical steps reported on by the

kcat, SVEs and SDKIEs were examined for Phe89Ala using the same methodology as

presented in Chapter 2 and Chapter 3. The lack of SVE suggests that a chemistry step is

rate-determining, rather than substrate binding or product release as might be expected

considering the involvement of Phe89 in binding, and the observed kcat is reporting on a

chemical step. The lack of a SDKIE suggests that the dehydration step has now become

fully rate-determining, as opposed to WT ssIGPS, in which the rate-determining step at

75 °C is ring closure. Using the methodology defined in Chapter 2, the dehydration step

is isotope insensitive, and the ring closure is isotope sensitive. These findings indicate

that the Phe89Ala substitution affects both substrate binding and catalytic events

associated with the dehydration step.

4.4.2 Interaction Between β1α1 and β2α2 Loops through Arg54 and Asn90 is

Important for Catalysis

Our studies with Lys53 (Chapter 3) and Phe89 amino acid substitutions indicate

that these conserved β1α1 and β2α2 loop residues are important for the chemical steps of

ssIGPS catalysis. Considering the SCA-MD results demonstrating an important

interaction between these loops through Arg54 and Asn90, we were also interested in

how the amino acid interaction between these loops may impact ssIGPS function. This

interaction may be involved in modulating the role of the β1α1 and β2α2 loops during

catalysis.8 To probe these interactions further, we made amino acid substitutions at both

Arg54 and Asn90, and characterized the kinetics of the variant enzymes at 75 °C, the

 

 

99  

biologically relevant temperature for the thermophile S. sulfataricus.

Arg54Ala and Asn90Ala display kinetic parameters very similar to WT enzyme at

75 °C, although Arg54Ala ssIGPS shows a small increase to kcat (~1.8 fold) compared to

WT ssIGPS. Although the overall rates were similar, it is still possible that the amino

acid substitutions could impact the individual steps of catalysis. To test this concept,

SVEs and SDKIEs were also measured in order to assign the rate-determining step for

each variant. In WT ssIGPS, the rate-determining step at 75 °C is the ring closure,

evidenced by the substantial SDKIE that was attributed the transfer of a single proton in

proton inventory studies (Figure 2.3).13 Neither Arg54Ala nor Asn90Ala exhibit an SVE

at 75 °C, suggesting kcat is reporting on a chemical event for both of these enzymes. Both

variants display a loss in the SDKIE at 75 °C that is similar to the behavior of previous

amino acid substitutions (Lys53Arg and Glu51Gln). Akin to these variants, the Arg54Ala

and Asn90Ala variants are likely affecting kinetics of the dehydration step, such that this

step becomes slower than the ring closure step and more rate-determining.

The Arg54Ala and Asn90Ala variants show changes to their pH profile (Figure

4.2), with an increase in the ascending pKa (attributed to general base activity by Gln51)

compared to WT enzyme occurring in both variants. The Asn90Ala variant also displays

a loss in the descending limb of the pH profile, whose pKa was previously attributed to

the general acid of the dehydration step, Lys53, although the same behavior is not seen

for Arg54Ala enzyme. The pH results provide evidence that the interaction between the

two residues is important for the dehydration step of the reaction. The

Arg54Ala/Asn90Ala double substitution was also examined and exhibited activity similar

to Asn90Ala. This result is evidence that the interaction between Arg54 and Asn90 is

 

 

100  

Figure 4.2: pH profiles of WT, Arg54Ala, and Asn90Ala ssIGPS show changes to the activity of the general acid and base. pH profile for (a) WT (pKa1 5.6 ± 0.2, pKa2 8.7 ± 0.1) is different than (b) Arg54Ala (pKa1 6.5 ± 0.2, pKa2 8.5 ± 0.2) and (c) Asn90Ala (pKa1 7.3 ± 0.3, pKa2>>9 ) . The amino acid substitutions result in peturbations associated with the general acid and base in the dehydration step compared to WT enzyme.  

 

 

101  

Table 4.2: pKa values for WT, Arg54Ala, and Asn90Ala indicate that the loop interaction is important for general acid/base catalysis. Enzyme pKa1 pKa2

WT 5.6 ± 0.2 8.7 ± 0.1

Arg54Ala 6.5 ± 0.2 8.5 ± 0.2

Asn90Ala 7.3 ± 0.3 >9

 

 

102  

important for function; but that Asn90 is the more vital residue, and the enzyme can more

easily overcome disruptions to Arg54 compared to Asn90.

The Arg54Ala and Asn90Ala variants change the rate-determining step of

catalysis at 75 °C. Our previous studies have also indicated that the rate-determining step

is temperature dependent. Therefore, to gain more insight into the effects of loop

substitutions, we also tested the temperature dependence of these variants. At lower

temperatures, Arg54Ala ssIGPS displays activity similar to WT ssIGPS. However, the

Asn90Ala variant shows a more substantial decrease in kcat compared to WT enzyme at

lower temperatures.

For WT ssIGPS, the rate-determining step at 75 oC is the ring closure; however, at

lower temperatures (25 oC) product release becomes rate-determining.13 At 37 °C both

the ring closure and product release contribute to the kinetic constants, so both are

partially rate-determining. The Asn90Ala variant shows no substantial SVE at 37 oC in

contrast to WT ssIGPS (Table 4.1). This finding suggests that kcat for the Asn90Ala

variant likely reflects a chemical step, and the effect of the Asn90Ala substitution on

chemistry is underestimated at the lower temperatures (i.e. for WT ssIGPS, kchemistry >>

kcat at 37 °C). Interestingly, there is still a significant SDKIE at 37 °C, as opposed to 75

°C at which both Asn90Ala and Arg54Ala enzymes display a loss in isotope effects. This

result indicates that the rate-determining step for these mutants is ring closure at 37 °C, as

opposed to 75 °C at which the dehydration step becomes rate-determining. For

Asn90Ala, the loss in SVE at 37 °C along with a significant SDKIE at this temperature

indicates that Asn90 is not only involved in dehydration, but also affects the rate of the

ring closure step, at least at lower temperatures.

 

 

103  

4.4.3 Analysis of Arg54Lys and Asn90Gln Variants of ssIGPS

Since Arg54 and Asn90 are residues that coevolve between IGPS enzymes from

different organisms, we were interested in examining amino acid substitutions that

mimicked this process. In the mesophilic enzyme ecIGPS, the analogous residues to

Arg54 and Asn90 are Lys55 and Gln94, respectively. These residue changes keep

hydrogen bond interactions intact at these positions. Arg54Lys and Asn90Gln amino acid

substitutions of ssIGPS were examined in order to further evaluate the role of this

interaction. These substitutions are expected to make similar interactions with CdRP;

however, they may be less optimal than the residues in WT due to differences in size

(Lys is slightly smaller than Arg and Gln is slightly larger than Asn).

The Arg54Lys variant displays activity similar to WT IGPS (Table 4.2), which is

expected considering the more deleterious amino acid substitution at this position,

Arg54Ala, also showed similar kinetic parameters to the WT enzyme. Despite the

similarity in kcat, between WT ssIGPS and the Arg54Lys variant, there is a substantial

difference in the magnitude of the SDKIE compared to WT enzyme at 75 °C but not 37

°C. This finding siggests that there is a difference in the step of the chemical mechanism

that is affected at lower versus higher temperatures. At the mesophilic relevant

temperature, 37 °C, the rate-determining step of the reaction is the ring closure, similar to

WT ssIGPS. Conversely, at the thermophilic-relevant temperature, 75 °C, the Arg54Lys

substitution causes the dehydration step to become slowed, making it the rate-

determining step at this temperature.

The Asn90Gln ssIGPS variant showed catalytic activity more comparable to WT

enzyme than those of the Asn90Ala variant, although the kcat was slightly reduced at 37

 

 

104  

°C compared to WT enzyme. Similar to the other substitutions evaluated, the SDKIE on

kcat was also substantially reduced compared to WT ssIGPS at 75 °C, indicating that the

Asn90Gln amino acid substitution also affects the dehydration step of the reaction,

despite the similar structure of Asn and Gln. At 37 °C, the SDKIE is present indicating

that the ring closure is rate-determining as shown previously. For completeness, the

double substitution, Arg54Lys/Asn90Gln ssIGPS, was also assayed. The kinetic

parameters are consistent with what was seen for the single substitutions, indicating that

the role of these residues are linked, and both are required for proper ssIGPS function.

4.4.4 Examination of the Interaction Between Coevolving Residues in the β2α2 Loop

In addition to the interaction between Arg54 and Asn90, there is a second active

site loop interaction identified using SCA-MD through Asn90 and Glu85, both of which

are located on the β2α2 loop. To further explore the role of this interaction, the Glu85Ala

variant was assayed. The steady-state kinetic parameters for Glu85Ala (Table 4.1) are

similar to WT ssIGPS. Interestingly, the KM at 75 °C is about six times higher than WT

enzyme, indicating that this residue may be involved in substrate binding. Similar to

Arg54Ala and Asn90Ala, Glu85Ala displayed a loss in the SDKIE at 75 °C, indicating

that this residue is also involved in the dehydration step of the reaction.

4.4.5 Structure and Stability of Loop Mutants

Amino acids residues like Arg54 and Asn90 may be important not only for

catalytic function, but may also for protein folding and stability, and changes to these

characteristics can also effect the activity of the enzyme.8 To assess the role of these

 

 

105  

amino acids on the structure and stability of ssIGPS, thermal inactivation experiments

were performed for WT and amino acid substituted ssIGPS. The thermal inactivation

experiments showed a similar rate constant of inactivation for WT and Arg54Ala ssIGPS

(1.11 x 10-3 s-1 and 1.24 x 10-3 s-1, respectively) (Figure 4.3). However, a two-fold

decrease in the rate of inactivation was observed for Asn90Ala (6.91 x 10-4 s-1) and

Asn90Gln (3.59 x 10-4 s-1) implying that changes to Asn90 increases the thermal stability

of the enzyme. The reason for this increase in thermal stability is unclear. The CD spectra

for WT and Asn90Ala ssIGPS (Figure 4.3) were nearly identical, indicating that the

substitution did not result in any gross structural change. The kinetic changes reported for

these variants must be then to local changes in the catalytic environment, likely through

changes in hydrogen bonding, rather than changes in structure and folding.

 

 

106  

Figure 4.3: Asn90Ala affects the thermal stability but not proper folding for ssIGPS. (a) Thermal inactivation curves show an increase in the thermal stability of the Asn90Ala variant compared to WT. (b) Circular dichroism curves indicate that the changes in activity for the Asn90Ala variant are not caused by gross structural changes to the enzyme and can be attributed to changes in the reaction.  

 

 

107  

4.5 Discussion

Lys53 and Phe89 are conserved and important amino acid residues for the IGPS

catalyzed reaction, and are found on the active site β1α1 and β2α2 loops, respectively.

The loss of the SDKIE for amino acid substitutions to Lys53 (presented in Chapter 3)

and Phe89 indicate that both residues are involved in the dehydration step of the reaction.

MD simulations previously identified correlated motion in these loops, which occurs

through a hydrogen bond between Asn90 and Arg54. The results presented in this

Chapter help to further delineate the role for the active site loops and the interaction

between them through Arg54 and Asn90 through the examination of enzymes with amino

acid substitutions at these positions.

The Asn90Ala variant showed a decrease in kcat at 37 °C compared to WT

enzyme, whereas at 75 °C, kcat approached that of WT enzyme. Conversely, the

Arg54Ala variant showed kcat values similar to WT ssIGPS at low temperatures, with a

slight increase at 75 °C. The changes to kcat for the variants were modest, but this does

not necessarily indicate that these residues do not play an important role in catalysis. The

SDKIEs suggest that the identities of the rate-determining step for WT and variant

enzymes are different, indicating that steady-state kcat values are underrepresenting the

true effects on the individual kinetic steps of the reaction. The substantial decreases in

SDKIEs for Arg54Ala and Asn90Ala variants suggest that the dehydration step is

becoming more rate-limiting relative to ring closure. Steady-state kinetics and careful

consideration of the SDKIEs and SVEs is likely the most efficient way of teasing these

effects apart, especially considering that pre-steady-state kinetics by the Goodey and

Sterner groups were unable to deconvolute the rate constants for the chemical steps of the

 

 

108  

reaction.

The temperature dependence of the Arg54Ala and Asn90Ala suggests that the

role of these residues may be affected by temperature. At 37 °C, there is still a substantial

SDKIE for both variants, indicating that ring closure is primarily rate-determining, as is

seen for the WT enzyme. This result further highlights the different temperature

dependencies of the chemical steps. We propose that at lower temperatures, the activation

energy for ring closure is much higher than the corresponding activation energy for

dehydration. At higher temperatures, the activation energies for the ring closure and the

dehydration steps are more similar. Upon the disruption of any amino acid involved in

dehydration, this step becomes rate-determining at 75oC, causing the loss of SDKIE seen

for the loop mutants. However, at lower temperatures, ring closure may still be primarily

rate-determining, leading to a less substantial effect on the SDKIE.

The decreased SDKIE for Arg54Ala and Asn90Ala variants along with the

changes to the pH profiles for these enzymes compared to WT ssIGPS indicates that

these coevolving residues effect the function of the general acid and base, Lys53 and

Glu51, in the dehydration step. The loss of the descending limb of the pH profile for

Asn90Ala is indicative of an increase in the pKa of the general acid, Lys53. Similarly, the

increase in pKa for the ascending limb of the pH profile for both Arg54Ala and Asn90Ala

variants suggests an effect on the general base, Glu51. Therefore, it is likely that Asn90 is

involved in proper function of Lys53, and affects the microenvironment and/or the

positioning of this general acid.

The results indicate that the interaction between the β1α1 and β2α2 loops helps to

properly bind the substrate and/or position Lys53 and Glu51 to act as the general acid and

 

 

109  

base of the dehydration step, and to increase catalytic efficiency by both allowing

residues that bind the substrate to make more favorable interactions. The lack of a similar

pH effect for the descending limb of the pH profile for the Arg54Ala variant can be

explained either by noting that the Asn90 side chain may hydrogen bond to the backbone

of Arg54, or to another residue on the β1α1 loop that is free to make an interaction (i.e.

Lys55 or Ser56) in the Arg54Ala variant. In this way, the Arg54Ala variant still

maintains the interaction between the β1α1 and β2α2 loops.

The interaction between the β1α1 and β2α2 loops between Asn90 and Arg54

functions to correctly position conserved amino acid residues, Glu51, Lys53 and Phe89.

The deleterious effect of the substitutions at these positions can be overcome at higher

temperatures due to the ability of the enzyme to more easily sample the conformation that

is required for proper catalysis by Glu51 and Lys53. This idea is in agreement with the

findings from the Bruice lab that showed the presence of NACs increases for ssIGPS as

temperature increases, and so the appropriate conformation for catalysis is more available

at higher temperatures.70, 78

This study provides further evidence that proper catalysis by IGPS requires the

reorientation of the substrate in the active site. We propose that at lower temperatures, the

enzyme has a slower rate of conformational sampling and lower thermal energy makes it

more difficult to rearrange the substrate after it is captured; thus, the ring closure is rate-

determining. Conversely, at higher temperatures, this reorientation is easier, decreasing

the activation energy for this step relative to the dehydration step.

The amino acid substitutions that mimic the analogous residues in ecIGPS,

Arg54Lys and Asn90Gln, display similar kinetic parameters to WT enzyme but have

 

 

110  

differences in the rate-determining step of the reaction at biologically relevant

temperatures. This finding indicates that the co-evolution of this loop and the particular

amino acids in these positions in ssIGPS are important to the kinetic mechanism of the

enzyme, and it is possible that these coevolving residues may play an important role in

the kinetic differences between the enzymes.

4.6 Conclusions

SCA-MD analysis provides additional insight into residue pairs that may not be

conserved but nonetheless play integral roles in enzyme catalysis. However, simply

identifying residues as important does not provide adequate information for application in

enzyme engineering, antibiotic development, or other industrial applications.

Experimental examination of the SCA-MD identified enzyme pairs provides a more in

depth understanding and additional details of the chemical mechanism that can be utilized

in other applications.

Our results provide an analysis of the interaction between the β1α1 and β2α2

loops through the SCA-MD identified Arg54 and Asn90. This interaction plays an

important role in the catalytic mechanism of ssIGPS and affects the role of Lys53 as a

general acid and Glu51 as a general base. Thus, it is likely that the interaction between

Asn90 and Arg54 is involved in guiding the anthranilate moiety of the substrate between

the two active sites and properly arranging the general acid for catalysis. These new

details can be used for the design of more efficient enzymes and perhaps new

antibacterial agents that target IGPS. Engineering new enzymes relies heavily in

understanding how a reaction occurs rather than just at what speed it occurs.

 

 

111  

4.7 References

1.  Hocker,  B.;  Jurgens,  C.;  Wilmanns,  M.;  Sterner,  R.,  Stability,  catalytic  versatility  and  evolution  of  the  (beta  alpha)(8)-­‐barrel  fold.  Current  Opinion  in  Biotechnology  2001,  12  (4),  376-­‐381.    2.  Carpenter,  R.  A.;  Xiong,  J.;  Robbins,  J.  M.;  Ellis,  H.  R.,  Functional  Role  of  a  Conserved  Arginine   Residue   Located   on   a   Mobile   Loop   of   Alkanesulfonate   Monooxygenase.  Biochemistry  2011,  50  (29),  6469-­‐6477.  3.  Malabanan,  M.  M.;  Amyes,  T.  L.;  Richard,  J.  P.,  A  role  for  flexible  loops  in  enzyme  catalysis.  Current  Opinion  in  Structural  Biology  2010,  20  (6),  702-­‐710.    4.  Lipchock,   J.;  Loria,   J.  P.,  Millisecond  dynamics   in  the  allosteric  enzyme  imidazole  glycerol   phosphate   synthase   (IGPS)   from   Thermotoga   maritima.   Journal   of  Biomolecular  Nmr  2009,  45  (1-­‐2),  73-­‐84.    5.  Rozovsky,  S.;  Jogl,  G.;  Tong,  L.;  McDermott,  A.  E.,  Solution-­‐state  NMR  investigations  of   triosephosphate   isomerase  active  site   loop  motion:  Ligand  release   in  relation  to  active  site  loop  dynamics.  Journal  of  Molecular  Biology  2001,  310  (1),  271-­‐280.    6.  Mazumder-­‐Shivakumar,   D.;   Bruice,   T.   C.,  Molecular   dynamics   studies   of   ground  state   and   intermediate   of   the   hyperthermophilic   indole-­‐3-­‐glycerol   phosphate  synthase.   Proceedings   of   the   National   Academy   of   Sciences   of   the   United   States   of  America  2004,  101  (40),  14379-­‐14384.    7.   Mazumder-­‐Shivakumar,   D.;   Kahn,   K.;   Bruice,   T.   C.,   Computational   study   of   the  ground   state   of   thermophilic   indole   glycerol   phosphate   synthase:   Structural  alterations   at   the   active   site   with   temperature.   Journal   of   the   American   Chemical  Society  2004,  126  (19),  5936-­‐5937.    8.  Shen,  H.  B.;  Xu,  F.;  Hu,  H.  R.;  Wang,  F.  F.;  Wu,  Q.;  Huang,  Q.;  Wang,  H.  H.,  Coevolving  residues   of   (beta/alpha)(8)-­‐barrel   proteins   play   roles   in   stabilizing   active   site  architecture  and  coordinating  protein  dynamics.  Journal  of  Structural  Biology  2008,  164  (3),  281-­‐292.    9.  Estabrook,  R.  A.;  Luo,  J.;  Purdy,  M.  M.;  Sharma,  V.;  Weakliem,  P.;  Bruice,  T.  C.;  Reich,  N.   O.,   Statistical   colevolution   analysis   and   molecular   dynamics:   Identification   of  amino   acid   pairs   essential   for   catalysis.   Proceedings   of   the   National   Academy   of  Sciences  of  the  United  States  of  America  2005,  102  (4),  994-­‐999.    10.  Dietrich,  S.;  Borst,  N.;  Schlee,  S.;  Schneider,  D.;  Janda,  J.  O.;  Sterner,  R.;  Merkl,  R.,  Experimental  Assessment  of   the   Importance  of  Amino  Acid  Positions   Identified  by  an   Entropy-­‐Based   Correlation   Analysis   of   Multiple-­‐Sequence   Alignments.  Biochemistry  2012,  51  (28),  5633-­‐5641.    

 

 

112  

11.   Hennig,   M.;   Darimont,   B.   D.;   Jansonius,   J.   N.;   Kirschner,   K.,   The   catalytic  mechanism  of  indole-­‐3-­‐glycerol  phosphate  synthase:  crystal  structures  of  complexes  of  the  enzyme  from  Sulfolobus  solfataricus  with  substrate  analogue,  substrate,  and  product.  J  Mol  Biol  2002,  319  (3),  757-­‐66.    12.  Schlee,  S.;  Dietrich,  S.;  Kurcon,  T.;  Delaney,  P.;  Goodey,  N.  M.;  Sterner,  R.,  Kinetic  Mechanism   of   Indole-­‐3-­‐glycerol   Phosphate   Synthase.   Biochemistry   2012,   52   (1),  132-­‐142.    13.   Zaccardi,   M.   J.;   Mannweiler,   O.;   Boehr,   D.   D.,   Differences   in   the   catalytic  mechanisms  of  mesophilic   and   thermophilic   indole-­‐3-­‐glycerol   phosphate   synthase  enzymes   at   their   adaptive   temperatures.   Biochemical   and   Biophysical   Research  Communications  2012,  418  (2),  324-­‐329.    14.   Hennig,   M.;   Darimont,   B.;   Sterner,   R.;   Kirschner,   K.;   Jansonius,   J.   N.,   2.0   A  structure   of   indole-­‐3-­‐glycerol   phosphate   synthase   from   the   hyperthermophile  Sulfolobus  solfataricus:  possible  determinants  of  protein  stability.  Structure  1995,  3  (12),  1295-­‐306.    

 

 

113  

Chapter 5 Conclusions

5.1 A New Understanding of Catalysis by IGPS The work presented in this dissertation focused on the kinetic and chemical

mechanism of ssIGPS, and has greatly improved the current understanding for this

enzyme. Previous to these studies, ssIGPS had only been extensively studied at lower

temperature (25 to 40 °C), which limits the biologically relevant information that can be

attained about catalysis since the enzyme is naturally found at higher temperatures.

Understanding the differences between thermophilic and mesophilic enzymes requires

rigorous analysis over a range of temperatures. The studies presented in Chapter 2

indicated that the identity of the rate-determining step of ssIGPS is temperature

dependent. At lower temperatures (25 °C), product release is rate-determining, but as

temperature increases to 75 °C, the ring closure step becomes rate-determining.

Interestingly, ecIGPS showed a different rate-determining step at its biologically relevant

temperature, indicating that differences in environment can affect the kinetic mechanism

of enzymes.1

Previous research also lacked a clear understanding of all conserved and

catalytically important active site residues. Some studies were at odds in the assignment

of the general base; crystal structures suggested Glu159 performed this role whereas MD

simulations implicated Glu210.2-4 There were also several conserved residues whose role

in catalysis was ambiguous including Glu51, Lys53, and Arg182. Lastly, the role of

Lys110 as the general acid in two different steps of the reaction is not typical in enzyme

catalysis and the mode of reprotonation was unknown. The comprehensive kinetic

 

 

114  

analysis on WT ssIGPS and variants for each of these conserved residues as presented in

Chapter 3 allowed a more complete understanding of the role of these residues. The

experiments unambiguously assigned Glu51 and Lys53 as the general base and acid

responsible for catalyzing the dehydration of the I2 intermediate to form the product. This

finding is at odds with previous suggestions that Glu159 (or Glu210) and Lys110

catalyze this step. However, this mechanism resolves the need to reprotonate Lys110

following the ring closure (where it donates a proton to the C2’ carbonyl). This

identification also indicates that there are two distinct active site surfaces, requiring that

the substrate undergo a reorientation in the active site after the formation of the I2

intermediate in order to place Lys53 and Glu51 in an appropriate position in order to

catalyze the dehydration.

In addition to analyzing those residues that are conserved, Chapter 4 examined

residues that are not conserved but are coevolving and showed correlated motion, both of

which can be important to the catalytic mechanism.5 Considering the importance of

active site loop residues Lys53 and Phe89, coevolving residues Arg54 and Asn90 were

examined. At 75 °C, the rate-determining step of the amino acid substitution Asn90Ala

was dehydration, in contrast to ring closure in WT ssIGPS. Additionally, the activity of

Asn90Ala variant was decreased compared to WT enzyme at lower temperatures,

indicating that this amino acid substitution is more deleterious at lower compared to

higher temperatures.

Similar evaluation of amino acid substitutions at the Arg54 position in ssIGPS

showed an important role for this residue as well. The results suggest that the interaction

between Asn90 and Arg54 is important for proper function of the general acid-base pair,

 

 

115  

Lys53-Glu51. Additionally, the deleterious effect of Asn90Ala variant is increased at

lower temperatures. This result along with MD simulations performed by Shen et al.

indicate that the interaction between the β1α1 and β2α2 loops through Arg54 and Asn90

may be involved in protein motions that are important for the chemical events of

catalysis.5 Together, these studies provide molecular level details on the role for active

site residues in ssIGPS catalysis, as well as a more complete view of catalysis by ssIGPS.

5.2 Implications for Understanding the Evolution of Thermophilic Versus Mesophilic Enzymes Efficient catalysis by enzymes requires a delicate balance between stability and

flexibility. Proteins from thermophilic organisms have developed an excess of stabilizing

interactions like salt bridges, in order to remain stable at high temperatures, which causes

a decrease in flexibility at lower temperatures. Conversely, their mesophilic homologs

lack these same structural elements, despite the similarity in structure and fold, allowing

increased flexibility at lower temperatures.6 The results from Chapter 2 shed some

insight into the evolutionary differences between thermophilic ssIGPS and mesophilic

ecIGPS. The two enzymes display differences in the rate-determining step of reaction. In

ssIGPS, the ring closure step is rate-determining, whereas this result is not seen for

ecIGPS.

This difference may be due to the added stabilizing interactions in ssIGPS that

create increased rigidity of the enzyme. In ecIGPS, there are fewer salt bridges, allowing

the enzyme to be more flexible. We propose that the ring closure step involves dynamic

motions in the enzyme that allow the bond between C1 and C2’ to form (as was also

proposed by Bruice and coworkers3, 4). ssIGPS cannot complete this step as quickly at

 

 

116  

lower temperatures due to limited flexibility, while it can occur more quickly in ecIGPS.

Therefore, in ssIGPS this step has become rate-determining. In agreement with this idea

is the temperature dependence of the kinetic mechanism for ssIGPS. The decreased

motion at lower temperatures slows this step more severely than at higher temperatures

causing changes to the kinetic mechanism at different temperatures.

5.3 Engineering New Indole Derivatives and Improving Industrial Indole Synthesis with IGPS Many different industries including pharmaceuticals, agriculture, and materials

would benefit from the integration of enzyme technology into their processes.7 As a

thermophilic enzyme, ssIGPS is robust and can remain stable under high temperatures

and harsh conditions, which makes it quite advantageous for industrial processes

compared to its mesophilic homologs. Additionally, the IGPS catalyzed reaction is

industrially relevant, as indole is a widely used structure in many applications.8 In the

typical industrial catalysis, indole and its derivatives are made using the Fischer indole

synthesis or the Japp-Klingemann reaction.9 While both of these reactions have been

optimized for the desired product, they require the use of high heat, extreme pH,

nonaqueous solvent, and heavy metal catalysts. Additionally, they require the production

of intermediates that can be difficult to attain, and they can create byproducts, which

decrease the efficiency of the reaction and make purification more difficult.9 Introduction

of IGPS for indole derivative production would resolve many of these issues, and allow

for more environmentally friendly processes that do not require the use of nonaqueous

solvents or heavy metal catalysts. The issue that remains with this application is the

introduction of a substrate derivative into the catalyzed reaction. The widely relevant

 

 

117  

indoles in industry are highly derivitized. Therefore, the active site of IGPS needs to be

adjusted to account for the new side chains.

The assignment of Lys53 and Glu51 as the general acid and base in the

dehydration step of the ssIGPS catalyzed reaction must be taken into consideration in the

design of enzymes for the industrial synthesis of indoles. Previous to this finding, the

unknown role for Glu51 may have resulted in its substitution in the design of an enzyme

for the production of an IGP derivative, which would prevent the proper catalysis by

IGPS. Similarly, side chains Arg182 and Glu210 are now known to be important for

substrate binding. The modification of the substrate may require that the position of these

residues be changed as well as the size of the side chain. For example, Arg182 may be

substituted with a Lys to accommodate a slightly larger substrate, or to Glu or Asp to

accommodate a positively charged group. Understanding how the WT residues impact

catalysis allows for more thoughtful design of a modified enzyme.

5.4 Improving Novel Enzyme Engineering Efforts

Despite the wide reaching applications of engineering novel enzymes capable of

catalyzing non-natural reactions, and the large amount of resources being used to

engineer such enzymes, scientists are still unable to match the rate enhancements

achievable by natural enzymes.10 Studies have shown that this may be due to a inability

to appropriately model the dynamics of the enzyme when designing the new enzyme

activity onto naturally occurring enzyme scaffolds; our understanding of enzyme

dynamics is still in its infancy.11 Bioinformatics approaches performed by Juritz et al.

examined the differences in dynamics of enzymes with similar tertiary structure and

 

 

118  

correlated the results with protein sequence.12 The results suggest that sequence diversity

evolved among enzymes to allow for differences in dynamics. This finding suggests that

there is a specific relationship between sequence and dynamics that has yet to be

sufficiently understood, but will be essential in expanding the application of enzymes in

industry. Our ability to understand how sequence diversity contributes to dynamics and

catalysis is essential to further the knowledge of the role for enzyme dynamics in

catalysis, but requires extensive analysis of how individual residues contribute to

catalysis and dynamics. Enzymes containing the (β/α)8-barrel fold have very similary

structures, but catalyze a diverse range of reactions, which makes this fold an excellent

model for understanding the relationship between sequence, structure, and dynamics.

The idea that the sequence diversity is related to dynamics among enzymes with

the same three-dimensional structure may help explain the coevolution of residues in

enzymes. Many times changing residues does not change interactions; however, these

changes may alter dynamic fluctuations in the protein. Dynamic fluctuations can also be

thought of as the making and breaking of noncovalent interactions like hydrogen

bonds.13, 14 In ssIGPS, the hydrogen bond interaction between Arg54 and Asn90 helps to

aid in the dehydration step of the reaction. In ecIGPS this same interaction is between

Lys55 and Gln94. The difference in the interaction, caused by differences in the residues

at these positions, may create differences in how the enzymes fluctuate, and may

represent one reason for the coevolution, especially considering that comparison of

ssIGPS and ecIGPS has shown inherent differences in their kinetic mechanisms.

It has been long understood that amino acid residues that do not participate

directly in chemistry can still be important for folding and other processes. The result that

 

 

119  

residues, like Arg54 and Asn90, can change the rate-determining step of the reaction

without showing significant changes in catalytic turnover also indicates that these

residues are important for the chemistry of the reaction. In enzyme engineering studies,

there is concern with understanding how a reaction proceeds. Based on the kinetic results

for the Asn90Ala mutant, it is clear that this variant affects proper function of the general

acid and base, Lys53 and Glu51. We propose that this amino acid is partially responsible

for the reorientation of the I2 intermediate in the active site. More specifically, Asn90Ala

and its interaction with the β1α1 loop, is involved in a conformational change that allows

for proper reorientation of the I2 intermediate prior to dehydration. The temperature

dependence of Asn90Ala ssIGPS activity indicates that its function is likely gated

through a dynamic process. Changing this residue (or other similar residues) in an

enzyme-engineering scaffold can create unexpected dynamic changes that can affect the

activity of the new model. Current algorithms for the design of enzymes are not able to

account for all of the dynamic changes in the enzyme that are caused by sequence since

we do not fully understand how sequence affects dynamics. This effect is perhaps part of

the reason current efforts toward engineering novel enzymes are falling short of natural

enzymatic rate enhancements. Studies like that presented here, along with experimental

studies of the enzymes dynamics in IGPS and other enzymes will help to develop this

understanding.

 

 

120  

5.5 Future Studies

This research has evaluated the roles for both conserved and nonconserved

residues in ssIGPS catalysis. One of the most intriguing findings from this work is that

the substrate must undergo a reorientation during the reaction, which may be aided by the

enzyme. The next step is to understand how the enzyme undergoes dynamic fluctuation

in order to aid in this step of catalysis, especially considering the results by Goodey and

Sterner implicate dynamics of the β1α1 loop in catalysis even at low temperatures (25

°C).15 In these studies, the role of conformational motion in the active site β1α1 loop was

examined using the addition of fluorescent dyes. The authors suggest that the binding of

substrate and the chemical steps of catalysis are accompanied by conformational changes

in the enzyme.16

To further evaluate the role of dynamics, and gain additional site-specific

information about the conformational sampling for the IGPS enzyme, NMR can be

utilized. NMR is a robust technique and allows enzyme dynamics to be probed across

multiple timescales ranging from picosecond to second. Additionally, it allows the

measurement of parameters for almost every amino acid in an enzyme, providing site-

specific resolution of dynamics and conformational fluctuation.16 Preliminary NMR

experiments utilizing Carr-Purcell-Meiboom-Gill (CPMG) R2 relaxation dispersion

indicated that ssIGPS does not undergo very much conformational exchange on the

microsecond to millisecond timescale at the temperatures assayed (293 to 313K). The

lack of conformational exchange in the relaxation dispersion experiments can be

explained several ways. First, it is possible that the motions that contribute to IGPS

catalysis are occurring on a different timescale, and thus are not able to be measured

 

 

121  

using this technique. Based on the kinetic results of WT and amino acid substituted

ssIGPS, we believe that loop motions help to reorient the substrate in the active site

during the reaction, but these motions may be too slow to be measured by relaxation

dispersion. Other NMR techniques that provide dynamic information on slower

timescales such as hydrogen-deuterium exchange spectroscopy may provide more

applicable information.

A second explanation for the lack of measurable conformational exchange by

relaxation dispersion is that these experiments were completed at low temperatures (20 to

40 °C) compared to the biologically relevant temperature of ssIGPS. Considering

previous results that indicate flexibility is decreased at lower temperatures and the

enzyme is more rigid, it is likely that even if the applicable conformational exchange of

IGPS is on the appropriate timescale, at lower temperatures the enzyme is too rigid and is

not undergoing the same exchange as at higher temperatures. This idea is consistent with

the results of Bruice et al. in which the number of NACs is lower at lower temperatures.3,

4 Unfortunately, the use of the cryoprobe in the NMR experiments prevents us from

performing the experiments at a temperature more biologically relevant for ssIGPS.

In order to overcome this issue, studying a mesophilic homolog like ecIGPS

would allow for NMR data to be collected at a temperature biologically appropriate for

the enzyme activity. However, NMR requires a large amount of protein compared to

traditional biochemical techniques, and the current ecIGPS construct in use in our

laboratory exhibits precipitation at high concentrations (~1 mM). The instability of the

enzyme may be caused by the truncation of the original gene, which contained both

ecIGPS and the preceding enzyme in the pathway, PRAI, which are found as one

 

 

122  

bifunctional complex in E. coli naturally. Use of the entire bifunctional complex would

allow for the higher concentrations needed for NMR without instability problems, but

this enzyme is large by NMR standards (~50 kDa). To make these studies more feasible,

methodologies similar to those used in our laboratory to study RNA-dependent RNA

polymerase from poliovirus17 be used to examine the ecPRAI:IGPS complex. In these

experiments only certain residues in the enzyme are labeled (e.g. methionine), rather than

the entire backbone, and these resonances are used as probes for examining enzyme

dynamics and function. This will decrease spectral crowding and allow for the dynamics

of the entire complex be examined, as they may be different from the monofunctional

ecIGPS.

In addition to understanding the dynamics of ssIGPS, NMR can also allow the

orientation of the substrate in the active site to be probed. While crystal structures of

IGPS in complex with both rCdRP and IGP have been produced, it is clear from the

kinetic analysis that these structures likely do not represent the catalytically relevant

complex, as the C1 and C2’ carbons in rCdRP are located too far apart for a bond to

form, and IGP is bound such that Lys53 and Glu51 are not appropriately positioned for

dehydration of I2 to IGP. Exchange-transferred nuclear Overhauser effect (tr-NOE)

experiments can be explored as they allow for the measurement of inter-nuclear distances

in the ligand bound to substrate. Additionally, tr-NOE experiments do not require the use

of a cryoprobe, and thus experiments can be performed at temperatures more biologically

relevant to ssIGPS. These studies may provide valuable information about the orientation

of substrate at biologically relevant temperatures.

The relationship between sequence and dynamics will be important for the future

 

 

123  

development of enzyme technologies. The kinetic analysis presented in this dissertation

provides the understanding for how amino acid substitutions in ssIGPS can affect the

activity, and the NMR studies on WT IGPS can provide insight into how motion

contribute to catalysis. In order to complete the understanding for how sequence in IGPS

or other (β/α)8-barrel enzymes is related to the dynamics, understanding how these

mutations change both the activity and the dynamics will complete the larger picture.

Therefore, utilization of NMR not only on WT enzyme but also on ssIGPS variants

should be explored.

5.6 Conclusions

This dissertation redefined the catalytic and kinetic mechanisms of the IGPS

enzyme. The results presented here will aid in a more complete picture of catalysis by

enzymes and will aid in the understanding of the relationship between sequence and

dynamics. It is also directly relevant to several different applications including enzyme

engineering and indole synthesis.

5.7 References

1.   Zaccardi,   M.   J.;   Mannweiler,   O.;   Boehr,   D.   D.,   Differences   in   the   catalytic  mechanisms  of  mesophilic   and   thermophilic   indole-­‐3-­‐glycerol   phosphate   synthase  enzymes   at   their   adaptive   temperatures.   Biochemical   and   Biophysical   Research  Communications  2012,  418  (2),  324-­‐329.    2.  Hennig,  M.;  Darimont,  B.  D.;  Jansonius,  J.  N.;  Kirschner,  K.,  The  catalytic  mechanism  of   indole-­‐3-­‐glycerol   phosphate   synthase:   crystal   structures   of   complexes   of   the  enzyme   from   Sulfolobus   solfataricus   with   substrate   analogue,   substrate,   and  product.  J  Mol  Biol  2002,  319  (3),  757-­‐66.    3.  Mazumder-­‐Shivakumar,   D.;   Bruice,   T.   C.,  Molecular   dynamics   studies   of   ground  state   and   intermediate   of   the   hyperthermophilic   indole-­‐3-­‐glycerol   phosphate  

 

 

124  

synthase.   Proceedings   of   the   National   Academy   of   Sciences   of   the   United   States   of  America  2004,  101  (40),  14379-­‐14384.    4.   Mazumder-­‐Shivakumar,   D.;   Kahn,   K.;   Bruice,   T.   C.,   Computational   study   of   the  ground   state   of   thermophilic   indole   glycerol   phosphate   synthase:   Structural  alterations   at   the   active   site   with   temperature.   Journal   of   the   American   Chemical  Society  2004,  126  (19),  5936-­‐5937.    5.  Shen,  H.  B.;  Xu,  F.;  Hu,  H.  R.;  Wang,  F.  F.;  Wu,  Q.;  Huang,  Q.;  Wang,  H.  H.,  Coevolving  residues   of   (beta/alpha)(8)-­‐barrel   proteins   play   roles   in   stabilizing   active   site  architecture  and  coordinating  protein  dynamics.  Journal  of  Structural  Biology  2008,  164  (3),  281-­‐292.    6.   Feller,   G.,   Protein   stability   and   enzyme   activity   at   extreme   biological  temperatures.  Journal  of  Physics-­Condensed  Matter  2010,  22  (32).    7.  Woodley,   J.  M.,  Protein  engineering  of  enzymes  for  process  applications.  Current  Opinion  in  Chemical  Biology  2013,  17  (2),  310-­‐316.    8.  Barden,  T.  C.,  Indoles:  Industrial,  agricultural  and  over-­‐the-­‐counter  uses.  Topics  in  Hetercyclic  Chemistry  2011,  26,  31-­‐46.    9.  Humphrey,  G.  R.;  Kuethe,  J.  T.,  Practical  methodologies  for  the  synthesis  of  indoles.  Chemical  Reviews  2006,  106  (7),  2875-­‐2911.    10.   Ruscio,   J.   Z.;   Kohn,   J.   E.;   Ball,   K.   A.;   Head-­‐Gordon,   T.,   The   Influence   of   Protein  Dynamics  on  the  Success  of  Computational  Enzyme  Design.  Journal  of  the  American  Chemical  Society  2009,  131  (39),  14111-­‐14115.    11.   Lassila,   J.   K.,   Conformational   diversity   and   computational   enzyme   design.  Current  Opinion  in  Chemical  Biology  2010,  14  (5),  676-­‐682.    12.   Juritz,   E.;   Palopoli,   N.;   Silvina   Fornasari,   M.;   Fernandez-­‐Alberti,   S.;   Parisi,   G.,  Protein   Conformational   Diversity   Modulates   Sequence   Divergence.   Molecular  Biology  and  Evolution  2013,  30  (1),  79-­‐87.    13.  Boehr,  D.  D.,  During  Transitions  Proteins  Make  Fleeting  Bonds.  Cell  2009,  139  (6),  1049-­‐1051.    14.   Hammes,   G.   G.,   Multiple   conformational   changes   in   enzyme   catalysis.  Biochemistry  2002,  41  (26),  8221-­‐8228.    15.  Schlee,  S.;  Dietrich,  S.;  Kurcon,  T.;  Delaney,  P.;  Goodey,  N.  M.;  Sterner,  R.,  Kinetic  Mechanism   of   Indole-­‐3-­‐glycerol   Phosphate   Synthase.   Biochemistry   2012,   52   (1),  132-­‐142.    

 

 

125  

16.  Mittermaier,  A.  K.;  Kay,  L.  E.,  Observing  biological  dynamics  at  atomic  resolution  using  NMR.  Trends  in  Biochemical  Sciences  2009,  34  (12),  601-­‐611.    17.  Yang,  X.;  Welch,  J.  L.;  Arnold,  J.  J.;  Boehr,  D.  D.,  Long-­‐Range  Interaction  Networks  in  the  Function  and  Fidelity  of  Poliovirus  RNA-­‐Dependent  RNA  Polymerase  Studied  by  Nuclear  Magnetic  Resonance.  Biochemistry  2010,  49  (43),  9361-­‐9371.    

 

 

126  

APPENDIX

Solvent Deuterium Kinetic Isotope Effect Analysis* *The following analysis was completed by Eric Yezdimer

To demonstrate the connection between the observed SDKIE and each discrete

mechanistic step, it is useful to consider the following generalized series of chemical

reactions as a feasible model for IGPS catalysis:

(A.1)

where S, ES, EI, EP, P, and E denote substrate, enzyme-substrate complex, enzyme-

intermediate complex, enzyme-product complex, product, and free enzyme, respectively.

Since the first intermediate is only thought to be fleetingly stable, and the release of

carbon dioxide is the first irreversible step of the reaction, the kinetic scheme was

simplified to only include one intermediate, the I2. The corresponding rate formulas for

each step are given by:

(A.2)

(A.3)

(A.4)

!

S + E"k#1

k1ES$

k2EI"

k#3

k3EP"

k#4

k4E + P

!

d[P]dt

= k4[EP] " k"4[E][P]

!

d[EP]dt

= k3[EI]+ k"4[E][P] " k"3[EP] " k4[EP]

!

d[EI]dt

= k2[EI]+ k"3[EP] " k3[EI]

 

 

127  

(A.5)

(A.6)

Under conditions where the initial substrate concentration is much larger than the initial

free enzyme concentration, [S]o>>[E]o, the following steady-state approximations for the

enzyme bound species can be applied:

(A.7) Using the mass balance equations for the enzyme,

(A.8) and substrate/product concentrations,

(A.9)

Equation A.7 can be rewritten into the familiar Michaelis-Menton form.

(A.10)

where k’cat and K’M are parameters defined by,

(A.11)

!

d[ES]dt

= k1[E][S] " k"1[ES] " k2[ES]

!

d[S]dt

= k"1[ES] " k1[E][S]

!

d[EP]dt

=d[EI]dt

=d[ES]dt

= 0

!

[E] = [E]o " [ES] " [EI] " [EP]

!

[S] " [S]o # [P]

!

d[P]dt

= k 'cat[E]o([S]o " [P])K 'M +[S]o " [P]

!

k'cat =k2k3k4

k3k4 + k2(k3 + k"3k4 )

 

 

128  

(A.12)

Within the limiting initial rate behavior of the Michaelis-Menton Equation (A.7), [P] is

assumed to be near zero, whichr reduces Equation A.12 to an extended Michaelis

constant,

(A.13)

K’M can be experimentally determined along with k’cat. Dividing equations A.11 and

A.13 provides an expression for catalytic efficiency, k’cat/K’M,

(A.14)

To understand the source of the experimental SDKIE for ssIGPS in the k’cat and K’M

constants, we assumed that ony rate constants for the chemical steps k2, k3, and k-3 are

potentially isotope sensitive. Both steps of IGPS catalysis involve proton transfers with

solvent exchangeable protons. Rate constants under H2O or D2O solvent conditions will

!

K 'M =(k2 + k"2)(k3k4 + k3k"4[P])k1(k2k3 + k2k4 + k3k4 + k2k"3)

!

K 'M =k3k4k2 + k"1k3k4

k1(k2k3 + k2k4 + k3k4 + k2k"3)

!

k 'catK 'M

=k1k2k3k4 (k2k3 + k2k4 + k3k4 + k2k"3)

(k2 + k"1)(k3k4 + k2(k3 + k4 + k"3))(k3k4 + k3k"4[P]+ k"3k"4[P])

 

 

129  

be denoted by either “H” or “D” subscripts, respectively. The SDKIE for k’cat and K’M

are given by

(A.15)

(A.16)

This result illustrates the ability of the SDKIE for k’cat/K’M to report on steps after an

irreversible step. Furthermore, if k-1<<k2, as is predicted to occur for enzymes such as

ssIGPS with strong susbtrate binding affinity, then the SDKIE for k’cat/K’M is reduced to

one.

It is also valuable to consider k’cat,H/k’cat,D under various limiting conditions in

order to highlight other factors that could suppress the SDKIE. In cases where the

conversion of ES to EI represents a much slower catalytic step than the conversion of EI

to EP (k2<<k3), the SDKIE for k’cat simplifies to

(A.17)

!

k'cat,Hk'cat,D

=k2,Hk3,H (k2,Dk3,D + k2,Dk4 + k3,Dk4 + k2,Dk"3,D )

k2,Dk3,D (k2,Hk3,H + k2,Hk4 + k2,Hk"3,H )

!

k 'cat,HK 'M ,H

k 'cat,DK 'M ,D

=k2,Hk3,H (k2,D + k"1)(k3,Dk4 + k3,Dk"4[P]+ k"3,Dk"4[P])k2,Dk3,D (k2,H + k"1)(k3,Hk4 + k3,Hk"4[P]+ k"3,Hk"4[P])

!

k'cat,Hk'cat,D

=k2,H (k2,D + k4 )k2,D + k4

 

 

130  

Under these conditions, the SDKIE should report on the conversion of ES to EI provided

that product release is similar to or faster than the irreversible formation of the

intermediate. If product release is very slow (k4<<k2<<k3), the SDKIE will reduce to one.

In the opposite situation where the conversion from EI to EP is much slower than

the conversion of ES to EI (k3<<k2), the SDKIE for k’cat is given by,

(A.18)

If product release is very slow under these conditions (k4<<k3<<k2)

(A.19)

This analysis indicates that under conditions of very slow product release (k4<<k2,k3), as

is predicted from SVE experiments for ssIGPS at 25 °C,1 the SDKIE for k’cat should

approach one if

(i) k2 << k3

Or

(ii) k3 << k2, and k-3 << k3

Under conditions of fast product release, as is predicted from SVE experiments for

ssIGPS at 75 °C, the SDKIE for k’cat should approach one if

!

k'cat,Hk'cat,D

=k3,H (k3,D + k4 + k"3,D )k3,D (k3,H + k4 + k"3,H )

!

k'cat,Hk'cat,D

=k3,H (k3,D + k"3,D )k3,D (k3,H + k"3,H )

 

 

131  

(i) k2 << k3 and k2 is isotope insensitive

Or

(ii) k3 << k2 and k3, k-3 are isotope insensitive.

To differentiate these two scenaries, the effects of amino acid substitutions on

residues predicted to be important for these steps (i.e. Glu51 and Lys53) must be

examined. Amino acid substitutions are predicted to substantially slow the second step

chemical step such that k3<<k2. Considering that Glu51Gln and Lys53Arg substitutions

result in an SDKIE of approximately one, these results suggest that k3 and k-3 are solvent

isotope insensitive. It follows that the observation of an SDKIE for k’cat for WT at 75 °C,

k2<<k3 as equation A.18 will also reduce to one if k3, k-3 are not isotope sensitive. This

logic is also consistent with proton inventory studies on WT ssIGPS that indicate a single

proton transfer event if responsible for the SDKIE.

 

 

  VITA

Margot J. Zaccardi

Education 2008-2013 Ph.D. in Chemistry The Pennsylvania State University, University Park, PA 2004-2008 B.S. in Chemistry, Research Option The Florida Institute of Technology, Melbourne, FL Publications Margot J. Zaccardi, Yezdimer O., Boehr, D.D. Functional identification of the general acid and base in the dehydration step of indole-3-glycerol phosphate synthase catalysis. JBC 2013, In Press. Margot J. Zaccardi, Mannweiler, O., Boehr, D.D. Differences in the catalytic mechanisms of mesophilic and thermophilic indole-3-glycerol phosphate synthase enzymes at their adaptive temperatures. BBRC. 2012, 418, 324-329. Margot J. Zaccardi, Olson, J.A., and Winkelmann, K. Preparation of Electrochemically Etched Tips for Ambient Instructional Scanning Tunneling Microscopy. J. Chem. Ed. 2010, 87(3), 303-310. Presentations Margot J. Zaccardi, Yezdimer, E.M., Boehr, D.D. “A new understanding of the catalytic mechanism of the tryptophan biosynthetic enzyme indole-3-glycerol phosphate synthase” Poster Presenter at the Enzyme Mechanisms Conference. San Diego, CA, January 3-7, 2013. Margot J. Zaccardi, Yezdimer, E.M., Boehr, D.D. “Understanding the catalytic function of protein dynamics in indole-3-glycerol phosphate synthase” Poster Presenter at American Chemical Society National Meeting. Philadelphia, PA, August 19-23, 2012. Margot J. Zaccardi, Axe, J.M., Boehr, D.D. “Mutational Analysis of Coevolving Residues in the Tryptophan Biosynthetic Enzyme Indole-3-glycerol Phosphate Synthase” Poster Presenter at Gordon Research Conference: Enzymes, Coenzymes, and Metabolic Pathways. Waterville Valley, NH, July 10-15, 2011.