ROLES OF LARGE LOOPS IN CATALYTIC VERSATILITY OF CATALASE-
PEROXIDASES: SIGNIFICANCE OF PERIPHERAL STRUCTURES IN 
IMPROVISING ENZYME FUNCTIONS 
 
by 
 
Shalley Nidhish Kudalkar 
 
 
 
 
A dissertation submitted to the Graduate Faculty of 
Auburn University 
in partial fulfillment of the 
requirements for the Degree of 
Doctor of Philosophy 
 
Auburn, Alabama 
May 7, 2012 
 
 
 
 
Copyright 2012 by Shalley Nidhish Kudalkar 
 
 
Approved by 
 
Douglas C. Goodwin, Chair, Associate Professor of Chemistry and Biochemistry 
Holly R. Ellis, Associate Professor of Chemistry and Biochemistry 
Christian R. Goldsmith, Assistant Professor of Chemistry and Biochemistry 
Christopher J. Easley, Assistant Professor of Chemistry and Biochemistry 
 
ii 
  
Abstract 
 
 
Catalase-peroxidases are bifunctional enzymes that catalyze the removal of hydrogen 
peroxide by two distinct pathways (catalase and peroxidase). They are central to antibiotic 
resistance in Mycobacterium tuberculosis and may be virulence factors in several dangerous 
human pathogens. These enzymes also hold much promise for engineering new enzymes to 
combat long-standing problems (e.g., environmental contamination by toxic pollutants).  
Currently, understanding of catalase-peroxidase structure and function is lacking to facilitate new 
drug development and enzyme engineering. The purpose of the research described in this 
dissertation is to understand the molecular basis for the unique catalytic abilities of catalase-
peroxidases to fully capitalize on their potential. The versatile catalytic abilities of these enzymes 
arise from an active site that is normally restricted to one activity (i.e., peroxidase). This is 
facilitated by three structures which are quite distant from the active site: a C-terminal domain 
and two large loops (LL1 and LL2). Given that catalase-peroxidases have two large loops absent 
from their fellow peroxidases, it is reasonable to suggest that these loops may serve to fine tune 
the active site for bifunctionality. Indeed, deletion of any of these structures has a substantial 
impact on catalase-peroxidase active site environment and catalytic function (e.g., selective loss 
of catalase activity). The main emphasis of this dissertation is to explore the roles of these two 
 
iii 
  
large loops i)  by identifying additional roles of LL1 in the unique catalytic properties of KatGs, 
and ii) by targeting the intersubunit interactions between LL2 apex and C-terminal domain. Site 
directed and deletion variants targeting LL2 apex and its length and various portions of LL1 
(Y226F KatG, KatG?209-228, KatG?200-214, KatG?LL1) were generated and investigated by heme 
environmental spectroscopy, steady-state and transient-state kinetic techniques. For LL1, all the 
variants including tyrosine showed a complete loss of catalase activity but the deletion variants 
showed substantial increase in peroxidase activity compared to wild type (wt KatG) and Y226F 
KatG. This increase in peroxidase activity was traceable to rapid reduction of high oxidation 
state intermediates by exogenous electron donors, and coincident prevention of enzyme 
inactivation by peroxides. The steady-state kinetic parameters of KatG?200-214 were comparable to 
wt KatG. In case of LL2, deletion variants showed near to complete loss of catalase activity with 
variable effects on peroxidase activity. The peroxidase activity with respect to H2O2 showed a 
substantial rate of turnover (even in excess of wt KatG) at sufficiently high H2O2 concentrations, 
along with two orders of magnitude lower apparent second order rate constants suggesting 
deficiency in the correct association of H2O2 in the active site leading towards it reduction to 
H2O. However, peroxidase kinetic parameters with respect to ABTS suggested that electron 
transfer to the heme by exogenous electron donors is unaffected and in some cases even 
enhanced. Concomitant with the shift in activity was a change in heme coordination from 
predominantly hexacoordinate high-spin to predominantly pentacoordinate high-spin. In contrast 
to deletion variants, the apex substitution variants of LL2 showed similar spectroscopic 
characteristics compared to wt KatG with little to no effect on the catalytic activities. The results 
of these studies emphasize the fact that both large loops despite of their peripheral locations 
 
iv 
  
serve to fine-tune the active site for its bifunctionality by acting as a gate keeper (LL1) to limit 
the typical peroxidase activity in favor of catalase activity and by maintaining the unique 
interaction occurring at 30? from the active site between LL2 and C-terminal domain. 
 
v 
  
Acknowledgments 
 
 
I would like to express my deepest gratitude to my advisor, Dr. Douglas Goodwin, for his 
excellent guidance, caring, patience, and providing me with an excellent atmosphere. His 
continual support and guidance throughout my research work made this dissertation a success. 
 I am thankful to my committee members Dr. Holly Ellis, Dr. Christian Goldsmith, and Dr. 
Christopher Easley, for their support and suggestions.  I am also thankful to Dr. Angela Calderon 
for agreeing to take on the role of the university reader.  I am grateful to Dr. Duin for all his help 
and advice with EPR.   
I would also like to extend my sincere thanks to Dr. Robert Moore, Elizabeth Ndonsta, Yu 
Wang, Haijun Duan, and Mick for their valuable suggestions and co-operation.  
My sincere thanks go to my best friend, Vinod, for his support and advice. I would like to 
thank my sisters- Meenakshi and Triveni, and brother- Paritosh for all their love and 
encouragement. I dedicate this work to my mother for her affection and prayers, to my daughter 
Nishqa, and to my husband, Nidhish, whose unconditional love and motivation made this 
research possible. 
 
 
vi 
  
Table of Contents 
 
 
Abstract ........................................................................................................................................... ii 
Acknowledgments........................................................................................................................... v 
List of Figures ................................................................................................................................ xi 
List of Tables ............................................................................................................................... xvi 
Chapter One: Literature Review ..................................................................................................... 1 
1.1. Heme and Heme Proteins......................................................................................................... 3 
1.2. Catalases .................................................................................................................................. 9 
1.3. Peroxidases ............................................................................................................................ 18 
1.3.1. Non-animal peroxidases .............................................................................................. 19 
1.3.2. Animal-peroxidases ..................................................................................................... 25 
1.4. Catalase-peroxidases .............................................................................................................. 29 
1.5. Conclusions ............................................................................................................................ 43
 
vii 
  
Chapter Two: Generation and Characterization of Catalase-peroxidases by Molecular and  
Biophysical Techniques .......................................................................................... 45 
2.1. Manipulating the katG gene: cloning and mutagenesis ......................................................... 45 
2.1.1. Site-directed mutagenesis ............................................................................................ 46 
2.1.2. Deletion mutagenesis ................................................................................................... 50 
2.2. Expression system .................................................................................................................. 51 
2.3. Protein purification ................................................................................................................ 53 
2.3.1. Soluble protein purification ......................................................................................... 53 
2.3.2. Insoluble protein purification ...................................................................................... 55 
2.4. Enzyme reconstitution ........................................................................................................... 55 
2.5. Spectroscopic characterization of KatG variants ................................................................... 56 
2.5.1. Circular dichroism spectroscopy ................................................................................. 57 
2.5.2. UV-visible absorption spectroscopy ............................................................................ 60 
2.5.3. Magnetic circular dichroism spectroscopy .................................................................. 67 
2.5.4. Electron paramagnetic resonance spectroscopy .......................................................... 70 
2.6. Kinetic characterization of KatG variants .............................................................................. 78 
2.6.1. Steady-state kinetic analysis ........................................................................................ 78 
2.6.2. Transient-state kinetic analysis of KatG ...................................................................... 81 
 
viii 
  
2.7. Conclusions ............................................................................................................................ 87 
Chapter Three: Tracing the Impact of a Unique Loop in Catalase-peroxidase Catalysis ............. 88 
3.1. Introduction ............................................................................................................................ 88 
3.2 Materials and Methods ............................................................................................................ 95 
3.2.1. Materials ...................................................................................................................... 95 
3.2.2  Site-directed and deletion mutagenesis ....................................................................... 95 
3.2.3. Expression and purification ......................................................................................... 97 
3.2.4. Absorption spectra and activity assays ........................................................................ 97 
3.2.5. Rapid freeze quench EPR spectroscopy ...................................................................... 98 
3.2.6. Stopped-flow analysis.................................................................................................. 99 
3.3 Results   ................................................................................................................... 100 
3.3.1. Steady-state kinetic analysis ...................................................................................... 101 
3.3.2. UV-visible characterization ....................................................................................... 106 
3.3.3. Transient-state kinetic analysis: ................................................................................. 108 
3.4. Discussion ............................................................................................................................ 127 
Chapter Four: Dependence of Catalytic Ability of Catalase-peroxidases on Intersubunit 
Interactions .......................................................................................................... 134 
 
ix 
  
4.1. Introduction .......................................................................................................................... 134 
4.2. Materials and Methods:........................................................................................................ 139 
4.2.1.Materials ..................................................................................................................... 139 
4.2.2. Cloning of Site-specific and deletion variants ........................................................... 140 
4.2.3. Expression and purification ....................................................................................... 142 
4.2.4. Circular dichroism spectroscopy ............................................................................... 142 
4.2.5. UV-visible absorption spectra and activity assays .................................................... 142 
4.2.6. Magnetic circular dichroism spectroscopy ................................................................ 143 
4.2.7. Electron Paramagnetic Resonance Spectroscopy and Spin Quantification ............... 144 
4.2.8. Evaluation of cyanide binding by titration and by stopped-flow .............................. 144 
4.3. Results .................................................................................................................................. 145 
4.3.1. Catalase activity ......................................................................................................... 149 
4.3.2. Peroxidase activity ..................................................................................................... 149 
4.3.3 Spectroscopic evaluation of KatG LL2 variants......................................................... 161 
4.3.4. Cyanide binding studies............................................................................................. 176 
4.4. Discussion ............................................................................................................................ 184 
 
x 
  
Chapter Five: Summary .............................................................................................................. 191 
5.1. Effect on KatG catalytic versatility: Residue conservation of Large Loop 1 ...................... 193 
5.2. Effect on KatG catalytic versatility: Length Conservation of Large Loop 2 ....................... 195 
5.3. Conclusions .......................................................................................................................... 197 
References?????????????????????????????????210
 
xi 
  
List of Figures 
 
 
Figure 1.1: Heme Structure ............................................................................................................. 5 
Figure 1.2: Various heme b ligand environments. .......................................................................... 7 
Figure 1.3: Catalytic cycle of catalases ......................................................................................... 11 
Figure 1.4: Structure of heme d .................................................................................................... 13 
Figure 1.5: Global structure of catalase. ....................................................................................... 14 
Figure 1.6: Active Site of Escherichia coli HPII catalase with important distal and proximal 
residues. ..................................................................................................................... 17 
Figure 1.7: Catalytic cycle of peroxidases .................................................................................... 24 
Figure 1.8: Structure of post-translationally modified and covalently linked heme b with the 
protein via ester bond formation with highly conserved aspartate and glutamate 
residues ...................................................................................................................... 26 
Figure 1.9: General reaction scheme of animal peroxidases ........................................................ 28 
Figure 1.10: Classical reaction scheme for catalase-peroxidase ................................................... 31 
Figure 1.11: Active sites of catalase-peroxidase (right) and monofunctional catalase (left). ....... 32 
Figure 1.12: Active Sites of catalase-peroxidase (right) and monofunctional peroxidase (left). . 34
 
xii 
  
Figure 1.13: C-terminal domain and two large loops of catalase-peroxidase. .............................. 36 
Figure 1.14: Model of (Trp-Try-Met) covalent adduct of catalase-peroxidase ............................ 40 
Figure 2.1: Overview of QuikChangeTM site-directed mutagenesis method. ............................... 47 
Figure 2.2: Overview of ?Around the horn? site-directed mutagenesis method. ......................... 49 
Figure 2.3: Far UV CD spectra associated with various types of secondary structures. .............. 59 
Figure 2.4: Typical absorption spectra for KatG in the soret (top) and charge transfer (lower) 
regions. ...................................................................................................................... 62 
Figure 2.5: Effects of ligands on FeIII d-electrons. ....................................................................... 64 
Figure 2.6: Further splitting of d-orbital energy levels in heme peroxidases. .............................. 65 
Figure 2.7: Electronic transitions observed in heme peroxidases. ................................................ 66 
Figure 2.8: Basis for an MCD spectrum: A-term. ........................................................................ 68 
Figure 2.9: Basis for an MCD spectrum: C-term. ......................................................................... 69 
Figure 2.10: Characteristic differences in ferrous MCD spectra of high-spin (black) and low-spin 
heme (gray). ............................................................................................................. 71 
Figure 2.11: Electronic paramagnetic absorption. ........................................................................ 73 
Figure 2.12: EPR absorption derivatives. ..................................................................................... 75 
Figure 2.13: Different EPR spectra observed in catalase-peroxidases ......................................... 76 
Figure 2.14: Basic setup for stopped-flow instrument. ................................................................. 83 
 
xiii 
  
Figure 2.15: Spectral changes (A and B) and typical time traces (C) observed for reaction of 
ferric enzyme with potassium cyanide. .................................................................... 85 
Figure 3.1: LL1 Connecting Distal and Proximal Part of the Active Site in KatG. ..................... 94 
Figure 3.2: Effect of H2O2 concentration on the peroxidase activity of wild-type KatG, Y226F 
KatG, and deletion variants (KatG?LL1, KatG?200-214, and KatG?209-228). ................ 104 
Figure 3.3: Effect of ABTS concentration on the peroxidase activities of the catalase negative 
variants KatG?209-228 (?), KatG?LL1 (?), and Y226F KatG (?). .............................. 107 
Figure 3.4: Effect of H2O2 on the absorbance spectra of Y226F KatG (A), and KatG?209228 (B).
 ................................................................................................................................ 109 
Figure 3.5: Compound I formation of Y226F KatG. .................................................................. 111 
Figure 3.6: Compound I formation of KatG?209-228. .................................................................... 112 
Figure 3.7: Rate constants for formation of compound I in Y226F KatG (?) and KatG?209-228 (?) 
plotted against H2O2 concentration at 400 nm. ....................................................... 113 
Figure 3.8: Compound I decay to compound II of Y226F KatG. ............................................... 114 
Figure 3.9: Compound I decay to compound II of KatG?209-228. ................................................ 115 
Figure 3.10: Rate constants for formation of compound II in Y226F KatG (?) and KatG?209-228 
(?) plotted against H2O2 concentration at 412 nm. ................................................ 117 
Figure 3.11: Typical time trace at 412 nm showing decay of compound I to compound II. ...... 119 
Figure 3.12: Heme intermediates observed during reactions with excess H2O2......................... 120 
Figure 3.13: EPR spectra of appearance of different species in KatG?209-228 during the reaction of 
resting enzyme with H2O2. ..................................................................................... 122 
Figure 3.14: EPR spectra of appearance of doublet species in Y226F KatG during the reaction of 
resting enzyme with H2O2. ..................................................................................... 123 
Figure 3.15: Reduction of compound II of Y226F KatG (A) and KatG?209-228 (B). ................... 125 
 
xiv 
  
Figure 3.16: Rate constants for reaction of Y226F KatG (?) and KatG?209-228 (?) to the ferric 
state plotted against ABTS concentration. ............................................................. 126 
Figure 3.17: Proposed intermediates for catalatic (inner pathway) and peroxidatic (outer 
pathway) activities of wild-type KatG. .................................................................. 131 
Figure 3.18: Proposed intermediates formed during the interrupted catalatic (inner dead end) and 
peroxidatic (outer pathway) activities of Y226F KatG and LL1 deletion variants.
 ................................................................................................................................ 132 
Figure 4.1: Position of LL2 with respect to the active site Heme............................................... 137 
Figure 4.2: Multiple sequence alignment of 20 selected representatives from Class I families with 
one sequence from a typical Class II representative. Presented here are the three 
large loops of KatG; LL1 (A), LL2 (B), and LL3 (C). ........................................... 147 
Figure 4.3: CD spectra of group I (A and B), group II (C), and group III (D) variants ............. 150 
Figure 4.4: Effect of H2O2 concentration on the catalase activity of wtKatG and group I variants.
 ................................................................................................................................. 153 
Figure 4.5: Effect of H2O2 concentration on the catalase activity of wtKatG, KatG?LL2 and group 
III variants ............................................................................................................... 154 
Figure 4.6: Effect of H2O2 concentration on the catalase activity of wtKatG and group II variants.
 ................................................................................................................................. 155 
Figure 4.7: Comparison of peroxidase activities of wtKatG and group I variants. .................... 158 
Figure 4.8: Comparison of peroxidase activities of wtKatG and group II variants. ................... 159 
Figure 4.9: Comparison of peroxidase activities of wtKatG and group III variants. ................. 160 
Figure 4.10: UV-Visible spectral comparison of ferric wtKatG and group I variants. .............. 162 
Figure 4.11: UV-Visible spectral comparison of ferric wtKatG with group II (A and B), and 
group III (C and D) variants. .................................................................................. 163 
Figure 4.12: Overlay of ferric MCD of wtKatG and group I variants. ....................................... 164 
Figure 4.13: Overlay of ferric MCD wtKatG, group II (A) and group III (B) variants. ............. 165 
 
xv 
  
Figure 4.14: Overlay of ferrous MCD spectra wtKatG and group I variants. ............................ 167 
Figure 4.15: Overlay of ferrous MCD comparison of wtKatG, and group II variants. .............. 168 
Figure 4.16: Overlay of ferrous MCD spectra of wtKatG, KatG?LL2 and group III variants. .... 169 
Figure 4.17: Comparison of EPR spectra of wtKatG and group I variants. ............................... 170 
Figure 4.17: Comparison of EPR spectra of wtKatG and group I variants. ............................... 171 
Figure 4.18: Comparison of EPR spectra of wtKatG and group II variants. .............................. 172 
Figure 4.19: Comparison of EPR spectra of wtKatG and group III variants. ............................. 173 
Figure 4.20: Titration of wtKatG (A), Q293A KatG (B), KatG?291 (C), and KatG?288-293 (D) with 
potassium cyanide .................................................................................................. 177 
Figure 4.21: Titration of wtKatG and group I, II, and III variants with potassium cyanide. ...... 178 
Figure 4.22: Cyanide binding studies of Q293A KatG (A), IEEQ-A KatG (B), and KatG?288-293.
 ................................................................................................................................ 181 
Figure 4.23: Pseudo-first-order-rate constants for the formation of cyanide complex of wtKatG 
and variants as a function of cyanide concentration. ............................................. 182
 
xvi 
  
List of Tables 
 
 
Table 3.1: Catalase kinetic parameters for wild-type KatG, Y226F KatG, and deletion variants
..................................................................................................................................................... 102 
Table 3.2: Peroxidase kinetic parameters with respect to H2O2 for wild-type KatG, Y226F KatG, 
and deletion variants .................................................................................................. 105 
Table 4.1: Classification of variants based on catalatic activity ................................................. 138 
Table 4.2: List of primers used for making site-directed and deletion LL2 variants.................. 141 
Table 4.3: Apparent catalase kinetic parameters ........................................................................ 151 
Table 4.4: Apparent peroxide-dependent peroxidase kinetic parameters ................................... 156 
Table 4.5: Apparent ABTS-dependent peroxidase kinetic parameters ....................................... 157 
Table 4.6: Ratios, expressed as percentages, of various EPR signal observed in wtKatG and 
variants ....................................................................................................................... 175 
Table 4.7: Dissociation constants for cyanide binding of wtKatG and the variants obtained from 
equilibrium titration studies ....................................................................................... 179 
Table 4.8: kon, koff, and KD for cyanide binding by wtKatG, and its LL2 variants ...................... 183
 
1 
  
Chapter One: Literature Review 
 
Enzymes are exceptional catalysts that are capable of astounding rate enhancement with 
great specificity under mild conditions. They catalyze reactions in enzymatic pathways with 
excellent chemo-, regio-, and stereo-selectivities. Naturally, there is substantial interest in 
harnessing these capabilities for reactions which lack suitable catalysts. Thus, there is a 
considerable interest in engineering new enzymes for novel applications demanded by, among 
others, the agricultural, food and pharmaceutical industries and also as a tool for molecular 
biology.  The problems which could be addressed with appropriately engineered enzymes are 
potentially global in scope (e.g., like contamination of the environment with unmanageable toxic 
waste, etc.), opening the possibility for substantial impact for humankind.  
Although the potential benefits to be achieved from enzyme engineering are substantial, 
the barriers to successful enzyme engineering are equally formidable. Primary among these 
obstacles is a lack of understanding of a highly complex structure-function relationship. 
Unlocking the secrets of this complex relationship can be approached in a number of ways, but it 
would seem that nature has mastered the art of manipulating structure to produce a broad panel 
of highly functional catalysts. Thus, a potentially fruitful strategy for enzyme engineering may 
be drawn from examples of the process accomplished naturally in biological systems.  
 
2 
  
  Here it is important to point out that successful enzyme engineering must depend not only 
on a coherent strategy, but also on a workable methodology. We would argue that at least in part, 
the failure of efforts in enzyme engineering have been due to flawed strategy. In other words, 
these approaches have represented great methodology but faulty strategy. Clearly, either one on 
its own is insufficient. If methods are applied without the right strategies, the fine balance which 
nature has achieved in enzyme engineering can be disrupted and consequently give rise to 
proteins that are unstable, cannot fold properly, and have no catalytic activity no matter how 
elegant the methodology may be. On the other hand, if the strategy is applied without appropriate 
methods success cannot be achieved on any reasonable time scale. Here we would part with 
nature as the methods employed there take far too long. 
Taken together, our approach is to use nature to inform strategy, but employ 
methodology provided by molecular biology to allow for pursuit of these strategies at a 
reasonable rate. A fundamental thread of the research presented in this dissertation is that 
catalase-peroxidases (also known to the field as KatG) represent an example where very clear 
elements of an enzyme engineering strategy can be discerned. Catalase-peroxidases are heme 
containing enzymes. These enzymes catalyze the removal of hydrogen peroxide by two distinct 
pathways: catalase and peroxidase. Unique structural features enable KatGs to carry out this 
bifunctionality using a single active site. The purpose of this literature review is to address 
questions like: What can be learned about the development of the catalytic properties of KatG?  
What does this elucidate in terms of a strategic approach that might be pursued? How might the 
function of KatG be manipulated?  
 
3 
  
This chapter will first establish how proteins containing the heme cofactor are rich 
resources for exploring and understanding the structure-function relationship. In order to 
understand this structure-function relationship various structural levels of heme proteins will be 
discussed in terms of their impact on the functions of these proteins. Generally, four structural 
levels dictate catalytic activity of heme proteins: the structure of heme cofactor itself, the 
characteristics of protein derived heme ligands, the non-ligand heme environment and the global 
structure of the protein. This chapter will detail the work done so far to understand these 
structural levels and will present some important examples which signify their role in heme 
protein functions. In doing so, it will become evident that voluminous data are present in 
literature, but they primarily support the first three structural levels with little dedicated to the 
fourth level (i.e., the contribution of the global structure of these proteins has received little 
attention). For this reason, various heme enzymes will be discussed with particular attention to 
their structure?function relationship. It is here that the importance of global structure of the 
protein (which includes unique structures peripheral and distant from the active site) in 
manipulating the catalytic activity of protein surfaces will be addressed. Through the examples 
provided in this chapter, the rationale for this research on roles of important peripheral structures 
in maintaining the active site architecture and consequently the catalytic versatility of catalase-
peroxidases will come into focus.   
1.1. Heme and Heme Proteins 
Heme is not only easily monitored by several spectroscopic techniques, but it is also 
highly versatile and widely used in nature as a cofactor. It is a prosthetic group that binds iron 
 
4 
  
within a tetrapyrrole, and although there is more than one type of heme used in biology, the 
diversity of function heme fills corresponds much more to the diversity of protein scaffolds 
which surround it rather than differences in covalent structure of the heme itself. Thus, heme is a 
cofactor used in oxygen storage and transport, substrate oxidation, ligand sensing, electron 
transfer, and other processes (1). As with any protein, heme-dependent proteins are usually 
classified into different superfamilies based on their structural and /or functional similarities. 
These superfamilies might have a primary function (oxygen carrier, substrate oxidation etc.), but 
a considerable functional overlap can be seen among them.  
The heme structure consists of a tetrapyrrole ring system complexed with iron. The four 
pyrrole rings are linked by methylene bridges giving rise to highly conjugated and roughly planar 
porphyrin. Usually, the heme iron is octahedrally coordinated by six heteroatoms. The four 
equatorial ligands are nitrogens from each of pyrrole and the remaining two axial ligands lie 
above and below the plane of the heme. Heme proteins collectively perform three main 
functions: catalysis of redox reactions using peroxides or oxygen plus externally supplied 
electrons (e.g., cytochrome P450, peroxidases, catalases, and catalase?peroxidases), storage 
and/or transport (e.g., myoglobin, and hemoglobin) and, electron transfer (e.g., cytochrome c) 
(2). The ability of this simple metalloorganic compound to perform diverse functions is dictated 
by four structural levels: heme structure itself, identity of protein derived heme-ligands, the non-
ligand protein environment and, finally the global structure of the protein.  
Although many forms of heme exist in nature, the three most abundant forms of heme are 
heme a, heme b, and heme c (Figure 1.1). Heme a and heme b differ at R1 and R3 positions. In  
 
5 
  
N
NN
N
F e
O H
O H
O
O
R
1
R
2
R
3
 
 
Heme a Heme b Heme c 
Chemical formula C49H56O6N4Fe C34H32O4N4Fe C34H36O4N4S2Fe 
Functional group at R1 hydroxyehtylfarnesyl -CH=CH2 -CH-CH3-SH 
Functional group at R2 -CH=CH2 -CH=CH2 -CH-CH3-SH 
Functional group at R3 -CH=O -CH3 -CH3 
 
 
Figure 1.1: Heme. Structures were taken from the PubChem Public Chemical Database using 
the CID numbers: heme a - 5288529, heme b - 4973, heme c ? 25202875 (3) 
 
 
 
6 
  
heme b they are vinyl and methyl groups, respectively, whereas hydroxyethylfarnesyl side chain 
occupies R1 and formyl group is present at R3 in heme a. These differences make heme a more 
hydrophobic and electron-withdrawing, increasing its reduction potential.  For this reason, heme 
a is found in mammalian cytochrome c oxidases, enzymes present at the end of electron transport 
chain (4).  Heme c differs from heme b by becoming covalently attached to the protein through 
two thioether linkages at R1 and R2. Due to these thioether bonds proteins containing heme c 
(e.g., cytochrome c) are chemically and thermally stable (5). The covalent linkage is suggested to 
provide structural support to the proteins as there is not much effect on the reduction potential of 
the proteins containing heme c (6).  One less commonly used form of heme exists in nature as 
heme d, it is found in Penicillium vitale catalase and E.coli catalase (7, 8).  
Heme b (protoporphyrin IX) is the most widely used in nature and is the prosthetic group 
found in all the heme enzymes discussed in this chapter. This kind of heme is found in 
hemoglobins, myoglobins, cytochrome P450s, and heme-based sensor proteins. Figure 1.2 shows 
various heme b ligand environments. For the heme proteins that do not directly bind an 
exogenous ligand, the axial ligands to the heme iron are supplied by protein itself. For example, 
in cytochrome b550 has two histidine ligands (9). By contrast, in heme proteins which do bind 
oxygen, hydrogen peroxide, or other small exogenous ligands, only one axial site (the so called 
?proximal side?) is occupied by a protein-based ligand. The so-called distal side of the heme is 
either unoccupied or a weakly associated water molecule. Histidine is widely used as the 
proximal ligand in heme proteins (e.g., hemoglobin, myoglobin) and heme peroxidases. 
Exceptions include cysteinate sulfur in P450s (10) and haloperoxidases (11), and phenolate 
oxygen of a tyrosine residue in heme catalases (12). Whether ligated on the proximal side by  
 
7 
  
 
H e mogl ob i n
C ytc h r ome  P 450
C atal as e
C ytoc h r ome  c  p e r oxi d as e
 
 
 
Figure 1.2: Various heme b ligand environments. Propionate groups of all structures are 
oriented towards right. Amino-acids color codes are as follows: Histidine - Blue, Phenylalanine/ 
Tyrosine ? Green, Tryptophan ? Magenta, Aspartate ? Pink, Arginine ? Dark yellow, Threonine 
? Orange, and Cystiene/Serine-yellow. 
 
 
 
8 
  
histidine, cysteine, or tyrosine, the distal side of the heme invariably forms part of catalytic site. 
Depending on the enzyme?s status in the catalytic cycle, the distal side can either be unoccupied 
or occupied, and if occupied is very often by an oxygen atom derived from H2O, O2 or peroxide.  
The non-ligand heme environment also influences the immediate active center 
environment. This includes the amino-acid residues not directly bound to the heme. These 
residues modulate the activity by maintaining the polarity of distal and proximal pocket of heme. 
They also regulate the accessibility of the substrate to heme center and heme edge by optimizing 
the access channel for the particular substrate. This non-ligand heme environment can provide 
new function to a protein. For example, haloperoxidases and cytochrome P450s both have 
cysteine as the proximal ligand but both catalyze very different reactions (13). Haloperoxidases 
mediate the oxidation of halides by hydrogen peroxide whereas cytochrome P450 catalyzes a 
monooxygenation reaction (13, 14). This difference is due to the way substrate channel is located 
in both the enzyme. In P450s, the edge of the heme is buried, whereas the heme iron center is 
accessible to the substrates making oxo-transfer a dominant reaction. In contrast, the heme edge 
of chloroperoxidases is more accessible allowing chlorination of number of aliphatic substrates 
(11).   
Up to now, it has been shown that the nature of heme, protein derived heme ligands, and 
non-ligand heme environment modulate the function of heme enzymes. However, the global 
structure of the proteins, comprising of the unique structures peripheral and distant to the active 
site, is simultaneously a crucial part of heme enzyme function and the least understood. The lack 
of exploration of this fourth structural level can be attributed to two main reasons.  First, it is 
difficult to specifically identify the important peripheral structures to target and how to address 
 
9 
  
them. Secondly, it?s been thought that this structural level is not nearly as important as the active 
site itself. However, years of synthesis and characterization of inorganic model complexes 
demonstrate, if nothing else that the whole protein is important. Also, nature and our own 
experience show that true change of function may not necessarily result from active site 
manipulation. Here, bifunctional catalase-peroxidase (KatG) is an ideal model to identify the 
unique distant structural features and help to explore their role in maintaining its bifunctionality. 
Also, it points out that nature regards and addresses such structures in devising new catalysts. 
 In terms of catalase-peroxidase function, there are three superfamilies that are 
particularly relevant: typical catalases, non-animal peroxidases, and animal peroxidases. In this 
chapter, these superfamilies will be discussed to better understand how the entire protein 
structure influences the heme environment to promote one function over the other. 
1.2. Catalases 
Catalases are ubiquitous enzymes. All aerobic organisms, whether prokaryotic or 
eukaryotic and even aero-tolerant anaerobes (e.g., the archeal Methanosarcina barkeri) (15) have 
at least one enzyme with catalatic activity. Catalases are organized into three main groups based 
on the variety of subunit sizes, different heme prosthetic groups, and the variety of sequence 
groups: the ?classic? monofunctional heme-containing enzymes, the catalase-peroxidases, and 
the non-heme or Mn-containing catalases. In addition, a diverse group of heme-containing 
proteins, such as chloroperoxidase, plant peroxidases, and myoglobin, exhibit low levels of 
 
10 
  
catalatic activity. This can be attributable to the presence of heme which alone can exhibit a very 
low catalase activity (16). 
Catalases catalyze decomposition of hydrogen peroxide (H2O2) to water and oxygen and 
the overall reaction can be simply represented as in reaction 1.1. 
2H2O2            2H2O + O2    (1.1) 
Catalases from all three groups follow essentially a two-step process for the decomposition of 
H2O2: H2O2 reduction followed by H2O2 oxidation. There are two distinct stages in the reaction 
pathway for the heme and non-heme catalases, however the mechanism involved in these two 
stages differ considerably for classic catalases versus that for non-heme catalases. This section 
will limit itself to a consideration of heme containing proteins with catalatic activity. In heme 
catalases, the first stage involves oxidation of heme iron using H2O2 as substrate in order to form 
a ferryl porphyrin ? cation radical intermediate (compound I) and H2O (16). In the second stage, 
compound I acts as two?electron oxidant, producing O2 and regenerating the ferric form the 
enzyme (Figure 1.3). The catalase activity is typically monitored by either one of  two common 
methods: monitoring the dioxygen evolved during catalysis using a Clark electrode (17) or 
monitoring the decrease in H2O2 by decrease in absorbance at 240 nm (18). 
In eukaryotes, catalases are present in the peroxisomes, where H2O2 is generated via 
oxidative processes (e.g., ?-oxidation of long chain fatty acids) (19). H2O2 is capable of initiating 
cellular damage by direct reaction with certain biomolecular components, but more often 
reactive species derived from H2O2 (e.g., hydroxyl radical) are the true damage-causing agent. 
Thus, catalases remove H2O2 and prevent the cells from encountering harmful reactive species. 
 
11 
  
 
 
 
 
 
 
 
 
 
 
Figure 1.3: Catalytic cycle of catalases 
 
 
 
 
F e I  I  I
H 2 O 2 H
2 O
F e I  V
H 2 O 2H
2 O  +  O 2
O
N a t i v e
C o m p o u n d  I
 
12 
  
More than 300 catalase sequences have been reported to date, and comparisons of these 
sequences reveal a strong sequence homology for about 400 amino acid residues (20, 21). Out of 
these, 12 residues are fully conserved and seven other residues differ in only one or two of the 
sequences. The distal histidine is one of the fully conserved residues as are the active site 
asparagine and the fifth ligand to the heme iron (i.e., the proximal tyrosine). Despite the 
sequence similarities catalases are divided into well-defined subgroups based on the polypeptide 
length: large subunit catalases (>700 residues per subunit) and small subunit catalases (<500 
residues per subunit). Hyderoperoxidase II (HPII) from E.coli is the largest of the heme catalases 
with 753 residues per subunit. A crystal structure has been solved for the enzyme, and it is used 
here as a representative for all catalase structures (20). Beside size, the small and large subunit 
catalases have two other characteristic differences. The first is that small subunit catalases 
contain heme b, which is modified to heme d in large subunit catalases (Figure 1.4), This heme is 
an oxidized protoporphyrin IX and is produced through the auto catalytic modification of the 
pyrrole ring opposite to the essential histidine producing a cis-hydroxychlorin ?-spirolactone. 
The second difference is that large subunit catalases carry an extra C-terminal domain with a 
flavodoxin topology whereas small subunit catalases can bind NADPH cofactor despite the 
absence of a nucleotide binding motif (22, 23).  
Monofunctional catalases are homotetramers, 200-340 kDa in size with four prosthetic 
heme groups. Each subunit contains four distinct structural regions: an amino-terminal domain, 
an anti-parallel eight-stranded ?-barrel domain, the domain connection and the ?-helical domain 
(Figure 1.5). In large subunit catalases, there is a fifth domain at the C-terminal end of the  
 
13 
  
 
 
N
N
N
N
F e
C H
3
H
3
C
O H
O H
O
O
O
O
O H
H O
O
O
 
 
Figure 1.4: Structure of heme d. Structure was taken from the PubChem Public Chemical 
Database using the CID number: heme d - 3581(3) 
 
 
 
14 
  
 
Figure 1.5: Global structure of catalase. Structure was taken from PDB accession domain 
1GGE. Different structures are color coded as follows: Amino-terminal arm ? Blue, 8-stranded 
?-parallel domain ? Green, Wrapping domain ? Cyan, ?-helical domain ? Magenta, Carboxy- 
terminal domain ? Orange, and  Extended hinge joining helical and carboxy domains ? Red. 
 
 
 
15 
  
protein. This accounts at least in part for the greater molecular weight of the subunits in these 
enzymes (7, 24-29). 
The N-terminal domain forms a hairpin structure and is involved in intersubunit 
interactions, the extent of which increases with the length of this amino-terminal- domain (24). 
This domain is buried between adjacent subunits and may serve to stabilize the homotetramers 
(25). The essential distal histidine is situated in C-terminus of the N-terminal domain. The 
position and reactivity of distal histidine is controlled by a hydrogen bond network formed 
between highly conserved residues from the ?-barrel domain. The anti-parallel ?-barrel domain 
is the largest domain, and it is a highly conserved part of the molecule. This domain consists of 
an eight-stranded antiparallel ?-barrel with six ??helical insertions in the turns between the 
strands. The first half of the domain contains amino-acid residues which define the distal side of 
the heme pocket, and in the small subunit enzymes, the second half continues to the NADPH 
binding pocket. 
The wrapping domain is an extended region of about 110 residues that connects the 
globular ?-barrel and ?-helical domains. Even though the domain has a low content of secondary 
structures, it contains ? -helix with the amino-acid residues which define the heme proximal side 
including the essential tyrosine. This domain participates in extensive intersubunit interactions 
particularly with residues from the amino-terminal-domain region from another subunit.  
The ?-helical domain contains four contiguous ?-helices and is not highly conserved. 
This domain binds NADPH. An extra C-terminal domain of about 150 residues has been seen in 
 
16 
  
in two large catalase, PVC (7) and HPII (26), structures. This domain contains high content of 
secondary structure elements organized with a ?flavodoxin-like? topology. 
Both the position of the heme and its environment are strongly conserved among 
catalases, including four amino acid residues which are considered essential for catalysis: the 
proximal tyrosine ligand, and histidine, asparagine, and serine all on the distal side of the heme 
(Figure 1.6). The serine is hydrogen bonded to N? of the histidine and is essential for maintaining 
the proper orientation and nucleophilic character of the immidazole, and thus stabilizes the distal 
heme pocket structure. The distal histidine is parallel to the plane of the heme and oriented above 
pyrrole ring. This unusual orientation seems to favor more extensive ? ? ? interactions between 
histidine and porphyrin presumably lowering the reactivity of compound I with reducing 
substrate (27). Conversely, in all other heme-containing proteins with a distal histidine, it is 
perpendicular to the plane of the heme causing increased reactivity of compound I with reducing 
substrates especially in peroxidases (27).  Interestingly, in the catalase distal histidine is 
absolutely required for the formation of heme d from protoheme b (28).  The coessential 
asparagine residue in the distal heme pocket is involved in compound I formation by promoting 
proton transfer to the histidine. This facilitates peroxide anion binding to heme and stabilizes the 
transition state for heterolytic cleavage of the O ? O bond (29). 
Catalases are very clearly more disposed toward two-electron reduction of compound I 
rather than two sequential one-electron steps via compound II. One feature of the catalase 
structure that contributed to this is a particularly large, narrow hydrophobic main substrate access 
channel at the lowest section which allows only diffusion of small molecules with low polarity.  
 
17 
  
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
Figure 1.6: Active Site of Escherichia coli HPII catalase with important distal and proximal 
residues. Amino acids are coded to following color scheme: Histidine - Blue, Tyrosine ? Green, 
Asparaginine ? Magenta, Serine-Yellow. 
 
 
 
 
18 
  
This limits the rate of one electron reduction of compound I by lowering the rate of 
diffusion of bulky organic substrates through the lower half of the access channel. Three 
channels have been shown to connect the active site heme with the surface in the crystal structure 
of CatF (12) and HPII variants (35). The main channel is perpendicular to the heme plane and 
provides the access for substrate H2O2 as shown by molecular modeling (36, 37). The second 
minor channel also known as lateral channel as it approaches heme laterally, almost in the plane 
of the heme and the third channel connects the heme with the central cavity. Not much evidence 
has been reported about the involvement of these two later channels in substrate transport to the 
active site heme. 
1.3. Peroxidases 
Peroxidases are ubiquitous in nature and use various peroxides as electron acceptors to 
catalyze a number of oxidative reactions. These peroxidases can be heme or non-heme proteins. 
Non-heme peroxidases are not evolutionarily linked and are classified into five independent 
families. The largest one is the thiol peroxidase which is divided into two different subfamilies 
(Glutathione peroxidases and Peroxiredoxines). The remaining four families include 
alkylhydroperoxidase, non-heme haloperoxidase, NADH peroxidase, and manganese catalase 
(38). The non-heme peroxidases are largely outside the scope of this dissertation, and thus will 
not be discussed in detail.  
Most heme-dependent peroxidases are classified into two large superfamilies: i) peroxidase-
cyclooxygenase and ii) non-animal peroxidases. As the name suggests, the non-animal 
 
19 
  
superfamily has representatives from fungal, plant and bacterial species (39, 40). Peroxidase-
cyclooxygenases are found primarily in vertebrate organisms (41), however, because the 
members of this superfamily were first identified in mammals, it is also referred to as the 
mammalian/animal peroxidases superfamily.  Interestingly, significant similarity in amino acid 
sequences have also been encountered in various invertebrates like arthropods, mollusks, 
crayfish, Caenorhabditis elegans and dorsophilla, so they are mainly identified as animal 
peroxidases (42).   
1.3.1. Non-animal peroxidases 
This superfamily arose from gene duplication of an ancestral gene and has been divided 
into three classes based on amino acid sequence alignment and similarity of biological function 
(39, 40). Class I contains intracellular peroxidases like  ascorbate peroxidase (APX) and  yeast 
cytochrome c peroxidase (CcP) (39, 43). Due to the generation of H2O2 as a byproduct of 
photosynthesis and respiration, APX plays a critical role in protection of chloroplasts of higher 
plants (44). As the name suggests, APX uses ascorbate as reducing substrate to facilitate  H2O2 
reduction to water (45). In contrast, CcP uses ferrocytochrome c to reduce toxic peroxide to 
water (39). The primary context of CcP is as mitochondrial enzyme where it confronts H2O2 
generated as a result of misfiring in the respiratory electron transport chain (46, 47). 
Class II of the non-animal superfamily is composed of secretory fungal peroxidases. This 
class includes lignin peroxidases (LiP) and manganese-dependent peroxidases (MnP), each 
named after one of its substrates , and Coprinus cinereus peroxidases (ARP or CiP) named after 
 
20 
  
its source black inkcap mushroom from Basidiomycete family (39). The main role of class II 
peroxidase is to degrade lignin in wood, and participate in secondary metabolism under 
conditions of limited supply of nutrients (48).  
Class III of the superfamily includes the secretory plant peroxidases, such as horseradish 
peroxidase (HRP-C), as well as soybean, barley, and peanut peroxidases. In contrast to class I , 
class III peroxidases are not primarily used for H2O2 detoxification, but rather are associated with 
a broad range of physiological processes in the plant life cycle.  These peroxidases are involved 
in phytohormone (e.g., indole-3-acetic acid) catabolism, wound healing, and cell wall 
metabolism (39, 45, 48). In addition, class III peroxidases can generate H2O2 and hydroxyl 
radical (ROS) in a controlled manner during the oxidative burst, hypersensitive response or cell 
elongation (49).   
Non-animal peroxidases are monomeric proteins with molecular weights ranging from 35 
to 45 kDa and contain heme b. Evaluation of typical non-animal peroxidase structures reveals 
two structural subdomains (proximal and distal) which appear to have arisen from an early gene 
duplication event. The major secondary structural component in the overall structure of non-
animal peroxidases is the ?-helix. There are 10 ?-helices (A?J) which are linked by loops and 
turns, and only a minor contribution from ?-structures in the overall peroxidase structure.  The 
heme group sits in the pocket formed by two anti-parallel helices, B on the distal side and F on 
the proximal side. Class I and II peroxidases posses additional helical groups like short helix B? 
between helices B and C which is absent in class III, however class III contains  three additional 
helices, D? , F? , F? (50). The proximal subdomain contributes proximal histidine ligand to the 
 
21 
  
heme iron and evolutionary conserved aspartate. Class I peroxidases lack any disulphide bridges, 
carbohydrate moieties and calcium ions. In contrast, class II and III enzymes are glycoproteins 
with four conserved disulphide bridges and two calcium ions. Although, the position of 
disulphide bridges and calcium ion binding sites differ in both classes of enzymes (39, 51), the 
presence of these features does provide a substantial stabilization of the active site conformation, 
particularly as it relates to the coordination state of the active site heme.   
Across the three classes of non-animal peroxidases there is a remarkable conservation of 
active site structure and catalytic residues. The active site of all non-animal peroxidases contains 
heme b prosthetic group ligated to the imidazole N? of the proximal histidine (52). A strong 
hydrogen bond is present between N?2 of the proximal histidine and conserved aspartate residue. 
This increases the anionic character of the histidine and is proposed to help peroxidases stabilize 
highly oxidized ferryl intermediates (52, 53). On the distal side, a strictly conserved histidine is 
positioned about 5 ? above the heme iron. This residue acts as a general base, serving to 
deprotonate H2O2 and is important for the formation of compound I. The distal side also contains 
a strictly conserved arginine which is also involved in compound I formation by acting as an 
electrostatic catalyst to offset a negative charge that develops across the O-O bond of the 
substrate H2O2 and facilitates the formation of H2O (54, 55).  
A tryptophan residue in class I peroxidases and phenylalanine in class II and III 
peroxidases (except Arthromyces ramosus peroxidases, which has leucine) is present in the distal 
and proximal side of heme (39, 56-58). In CcP of class I peroxidase the proximal tryptophan acts 
as a cation radical site of compound I (59, 60). The site of protein radical was confirmed to be 
 
22 
  
proximal tryptophan adjacent to proximal histidine by site-directed mutagenesis studies (61) and 
EPR studies of compound I with deuterium labeled tryptophan residues (60).  But in case of 
APX, porphyrin radical is observed even though APX and CcP share sequence and structural 
similarities. However, the crystal structure of APX shows a K+ ion in the proximal cavity of 
heme which is absent in CCP (62). The K+ ion increases the redox potential of the proximal 
tryptophan making it more difficult to oxidize. Consequently, in compound I of APX the free 
radical is present as a porphyrin radical rather than  tryptophan radical (63). In class II and class 
III peroxidases, tryptophan is replaced by phenylalanine. Because electron withdrawal from this 
side chain is far more difficult, the radical remains centered in the porphyrin ? electron system 
(57, 58).  
In class II peroxidases two calcium ions, one near the proximal phenylalanine or leucine 
and second at the distal side of the heme, are present (64). The distal calcium ion is required for 
activity because it maintains the structural integrity of class II peroxidases. In MnP loss of this 
calcium ion disrupts active site such that the distal histidine becomes sixth ligand to the heme 
iron and induces a shift in the spin state from high-spin (active) to low spin (inactive) (65). As 
with class II peroxidases, heme environment in class III peroxidases is also sensitive to the 
presence or absence of calcium ion but a much less dramatic effect on the activity in the absence 
of calcium ions has been observed (50, 66, 67).  
 According to the crystal structures obtained to date, the active site of peroxidases show 
the presence of at least four ordered water molecules, three in the distal pocket and one in the 
proximal one (51, 68). The proximal water molecule is hydrogen bonded to the proximal 
 
23 
  
aspartate. Though the exact hydrogen bond network differs between peroxidases, the general 
pattern of the distal cavity water molecules involves an extensive network of hydrogen bonds 
connecting the distal histidine, distal arginine and pyrrole IV propionate of the heme (51). One of 
the water molecules is situated right above the heme but is typically taken to be far away for the 
direct coordination. But in some peroxidases (e.g., in CcP0 it is ligated to heme at the sixth 
position giving rise to high-spin hexacoordinate heme (69). The week coordination of this distal 
water molecule at the sixth position of the heme has been shown to be very important in catalysis 
for some of the heme peroxidases (70). 
All heme peroxidases utilize heme iron to transfer electrons from a reducing substrate (RH) 
to H2O2 (reaction 1.2). The overall reaction can be written as: 
H2O2 + 2RH       2H2O+ 2R?   (1.2) 
Figure 1.7 shows the detailed catalytic cycle of a typical heme peroxidase (39). The first step 
is same as that of the catalases, which is the oxidation of resting ferric enzyme by one equivalent 
of H2O2 to generate H2O molecule and compound I.  Compound I is then reduced by one-
electron from a reducing substrate to generate one equivalent of substrate radical (R?) and 
compound II (oxo-ferryl species). Lastly, Compound II is reduced back to resting state with the 
concomitant one-electron transfer from the reducing substrate to generate another equivalent of 
substrate radical and one equivalent of H2O. The main difference in compound I and compound 
II is their heme states. Unlike compound I which contains a porphyrin ?-cation radical, 
compound II is a fully covalent ferryl-oxo heme intermediate (39). 
 
 
24 
  
 
F e
I I I
H
2
O
2
H
2
O
F e
I V
O
R H
R
F e
I V
O
R H
R
N a t i v e C o m p o u n d  I
C o m p o u n d  I I
H
2
O
2
H
2
O
F e
I I I
O
2
C o m p o u n d  I I I
H
2
O  +
 
(RH: reducing substrate)  
   
Figure 1.7: Catalytic cycle of peroxidases 
 
 
 
25 
  
In the presence of excess H2O2, compound III, a ferrous dioxy/ferric-superoxide inactive 
intermediate, can be formed by three different routes. Compound III is formed by the reaction of 
compound II with H2O2. Native peroxidase reaction with superoxide (O2.-) or its protonated form 
(HO2.) can produce it too. Finally, reaction of ferrous enzyme with O2 produces compound III 
(39). Compound III is an inactive species in monofunctional peroxidases and it?s a dead end for 
the peroxidase cycle.  In the absence of H2O2, the reversion of compound III is relatively slow 
and in the presence of excess of H2O2, the additional reaction of compound III leads to 
irreversible inactivation of the peroxidases (71, 72). However, the presence of efficient reducing 
substrates can prevent this irreversible inactivation (73, 74). The formation of compound III via 
later two routes is irreversible (75). 
1.3.2. Animal-peroxidases 
The animal and non-animal peroxidase superfamily have evolved independently, 
providing a good example of convergent evolution (42).  Animal peroxidases differ substantially 
in sequence, overall structure and nature of the prosthetic group compared to non-animal 
peroxidases. The most obvious structural difference between these two superfamilies is the 
nature of the prosthetic group. In contrast to the members of the non-animal superfamily that 
have noncovalently bound heme b at the active site, the heme group of peroxidases from 
mammalian superfamily is post-translationally modified to form ester bonds with the protein via 
highly conserved aspartate and glutamate residues (Figure 1.8) (76, 77). The most well known 
animal peroxidases are myeloperoxidase (MPO) and lactoperoxidase (LPO). These enzymes are 
important in human biology and contribute in host defense against pathogens (41, 78). They play  
 
26 
  
 
 
 
Figure 1.8: Structure of post-translationally modified and covalently linked heme b with 
the protein via ester bond formation with highly conserved aspartate and glutamate 
residues 
N
NN
N
F e
C H
C H
2
C H
3
C
H
2
H
3
C
H
2
C
O H
O H
O
O
O
C
O
C H
2
O
C
C H
2
O
C H
H N
H
C
O
C H
2
H C
C
O
N H
C
O
C H
H
2
C
H N
H
2
C
S
H
3
C
C H
M e t
G l u
A s p
 
27 
  
an important part in the innate immune system by oxidizing small ions like halides, 
pseudohalides, and nitrite to their corresponding oxidation products, for example, hypohalous 
acids which are used as part of a bactericidal response (41, 78, 79).  Figure 1.9 depicts the 
general halide oxidation scheme for animal peroxidases. 
MPO is a cationic 146 kDa dimer consisting of two symmetry-related halves (~73 kDa) 
linked covalently by a disulphide bridge (80). Each half contains two polypeptides, a small one 
composed of 106 amino acids (14.5 kDa), and large one of 467 amino acids (58.5 kDa), a heme 
prosthetic group, a calcium ion, and five sites of asparagine-linked glycosylation.  The secondary 
structure is dominated by 22 ?-helices which comprise 36% of the entire molecule (79, 81, 82).  
Four helices from the large polypeptide, H5, H6, H8, and H12 and one from the small 
polypeptide, H2, form a core unit containing the heme group. 
The active site of MPO contains one heme group which is derivative of 
ferriprotoporphyrin IX. The modification of methyl groups on pyrrole rings A and C allow the 
formation of ester linkages with the carboxyl group of glutamate of the large and aspartate of the 
small polypeptide. In addition, heme is covalently bound to the sulfur atom of a methionine 
residue via a sulfonium linkage (82). Histidine serves as a proximal ligand which forms a 
hydrogen bond with an asparagine, these two residues are conserved in all animal peroxidases. 
The modified heme is positioned at the bottom of a deep crevice which allows only H2O2 and 
small anions to access the iron atom (81). The formation of compound I in MPO is reversible 
requiring a larger excess of H2O2 for its formation than the typical peroxidases (39). For  
 
 
28 
  
 
 
 
 
 
 
(HOX: hypohalous acid; X-: halide) 
 
Figure 1.9: General reaction scheme of animal peroxidases 
 
 
 
 
F e I I I
H 2 O 2 H 2 O
F e I V O
N a t i v e C o m p o u n d  IH O X X -
 
29 
  
example, HRP is converted to its compound I state upon addition of just one equivalent of H2O2 
(39). The unique feature of MPO is the reaction of compound I with chloride ion to produce 
hypochlorite ion (83). The formation of this bacterial killing hypochlorite ion is the critical part 
of neutrophil response to bacterial invasion (39). 
Lactoperoxidase (LPO) also contains covalently liked protoporphyrin IX type heme. LPO is 
glycoprotein with a single chain that has a molecular mass of ~68 kDa. The LPO structure 
consists of 20 ?-helices, 2 small anti-parallel ?-strands, 4 glycosylated asparagines residues; 1 
phosphorylated serine, 7 disulphide bridges and a tightly bound Ca2+ ion (84). Lactoperoxidase 
also forms compound I upon reaction with H2O2, but only stochiometric amounts of H2O2 are 
needed (39). However unlike MPO, LPO compound I is unable to oxidize chloride ion, but is 
capable of oxidizing other halides (like iodine or bromine) and pseudohalides (e.g., thiocyanate) 
to the corresponding hypohalite ions, which are potent antimicrobials (85).  
1.4. Catalase-peroxidases 
The preceding discussion sufficiently reveals that out of the four levels of protein 
structure, first three (i.e., heme, protein-derived ligands, non-ligand heme environment) and their 
effects on a given protein?s functional capabilities, are bolstered by a wealth of information.  It 
has also become obvious that comparatively  little is known about the fourth level. That is, the 
mechanism by which distant protein structures influence the active site environment, and thus, 
contribute to the functional capabilities of heme in different proteins is poorly understood. For 
reasons to be described in this section, the catalase-peroxidases provide an excellent model to 
 
30 
  
evaluate this aspect of the protein structure and function. Moreover, the research described in this 
dissertation takes the advantage of nature?s strategies towards enzyme engineering by using 
catalase-peroxidases as a focal point to understand the mechanism by which distant protein 
structures control the heme environment and modulate the activities of heme proteins. 
Catalase-peroxidases (KatGs) are present in prokaryotes and fungi. These enzymes are 
capable of decomposing H2O2 via two distinct mechanisms using a single active site. Though it is 
an over simplification, the catalytic cycle of KatG can be described by super-positioning of both 
catalase and peroxidase catalytic cycles (Figure 1.10) (86-90). The two cycles are plainly 
suggested to be diverting at compound I. That is, compound I could either be reduced by 
hydrogen peroxide (catalase) or an exogenous electron donor (peroxidase). In the catalase cycle, 
compound I is reduced by using another equivalent of H2O2, forming H2O and O2. Whereas in 
peroxidase cycle, compound I undergoes two single-electron reduction steps by an exogenous 
electron donor. The first reduction produces a substrate radical and compound II. The second 
reductive step produces ferric enzyme, a second equivalent of substrate radical and one molecule 
of H2O. 
Despite using a single active site to carry out both the activities, KatGs exhibit a 
prominent catalases activity and a substantial peroxidase activity with broad specificity. This 
bifunctionality is unique because typically catalases are poor peroxidases and peroxidases are 
weak catalases. Thus, the bifunctionality of KatGs raises fundamental questions. The first is to 
which group of enzymes do KatGs belong, catalases or peroxidases. The answer can be sought 
by sequence and structural comparison of these groups of enzymes. The overall amino acid  
 
31 
  
 
 
(RH: reducing substrate, R?: substrate radical) 
          
Figure 1.10: Classical reaction scheme for catalase-peroxidase 
F e
I  I  I
H
2
O
2 H
2
O
F e
I  V
H
2
O
2H
2
O  +  O
2
O
R H
R
F e
I  V
O
R H
R
N a t i v e
C o m p o u n d  I
C o m p o u n d  I I
H
2
O  +
 
32 
  
 
 
 
 
 
Figure 1.11: Active sites of catalase-peroxidase (right) and monofunctional catalase (left). 
Amino acids are coded to following color scheme: Histidine - Blue, Tyrosine/Arginine ? Green, 
Asparaginine ? Magenta, Serine/Trytophan ? Yellow, Aspartate-deep salmon red. 
 
 
 
 
33 
  
sequence and structural fold of KatG bear no similarity to monofunctional catalases. Even the 
active site of both the groups bears little resemblance if any to one another (Figure 1.11). 
On one hand, monofunctional catalases use either heme b or heme d as the prosthetic 
group (29, 30); KatGs always have heme b in their active sites (91-94). Additionally, the 
proximal and distal active site residues are different in both the groups. On the proximal side of 
heme, the fifth ligand in catalases is tyrosine whereas in KatGs it is histidine. On the distal side, 
histidine is the only common residue in both groups of enzyme but its orientation differs in these 
enzymes. In KatGs, this distal histidine is perpendicular to heme whereas in monofunctional 
catalases it is parallel.  In both the cases, distal histidine allows the proper binding and reduction 
of peroxide molecule. However, the perpendicular orientation of this histidine in KatGs favors 
increase reactivity of compound I with reducing substrate.  On the other hand, the sequence and 
structural comparison of KatGs to monofunctional peroxidases shows that these two groups of 
enzymes are closely related. Indeed, the first three structural levels of these two important groups 
of enzymes are nearly indistinguishable. The active site of KatG is virtually identical to 
monofunctional peroxidases with identical distal and proximal catalytic triad of amino-acid 
residues (Figure 1.12) (91-94). On the proximal side of the heme, the essential histidine is the 
fifth coordination ligand to the heme and it forms hydrogen bond with fully conserved aspartate, 
which also interacts with the indole nitrogen atom from the proximal tryptophan. The distal side 
involves three catalytically important residues i.e., arginine-tryptophan-histidine. In KatG, the 
histidine-arginine pair is important for compound I formation and the distal tryptophan is 
essential for compound I reduction by H2O2 (95-97). 
 
34 
  
 
 
 
 
 
 
 
Figure 1.12: Active Sites of catalase-peroxidase (right) and monofunctional peroxidase 
(left). Amino acids are coded to following color scheme: Histidine - Blue, Arginine ? Green, 
Trytophan ? Yellow, Aspartate-deep salmon red. 
 
35 
  
The highly similar active sites and structural similarities of KatGs to monofunctional 
peroxidases raises the next essential question. What strategies has nature used to incorporate a 
new activity into an old scaffold? In other words, what enabled KatGs to utilize monofunctional 
peroxidase active site to carry out catalase activity? The quest to solve this question has led to 
understanding the role of peripheral structures of KatGs, which are away from the active site and 
are generally absent from the monofunctional peroxidases, in maintaining the heme environment 
of these enzymes to impart the bifunctionality. Solution to this question will not only help us 
appreciate the nature?s preference in utilizing the old scaffold for new activity over developing 
an entire new enzyme system to incorporate new functions. But, will also aid in deciphering the 
impact of the fourth structural level of a protein on its function.  
Comparisons of the global structures of these two groups of enzymes reveal that KatG 
contains two extended loops in its active-site bearing N-terminal domain and also carries an 
additional domain (C-terminal) derived from gene duplication and fusion (Figure 1.13). Most 
importantly, both extended loops and the C-terminal domain are peripheral to an distant from the 
active site. The overall structure of the C-terminal domain is similar to that of the N-terminal 
domain consistent with its origin in gene duplication and fusion (98, 99). Both the domains have 
high ?-helical composition with few ?-sheets (91-94). As in monofunctional peroxidases, there 
are 10 major ?-helices (A-J) in each domain. However, only the N-terminal domain contains the 
active site and binds heme. This active site is buried inside the N-terminal domain and substrate 
access to the heme is through a narrow access channel which allows small substrates but is too 
restricted for the passage of larger substrates (43). Even though the C-terminal domain maintains 
the overall protein architectures of the active site, it does not bind heme or catalyze any reaction  
 
36 
  
 
Figure 1.13: C-terminal domain and two large loops of catalase-peroxidase. Structure was 
taken from PDB accession domain 1SJ2. Different structures are color coded as follows Heme-
red, N-terminal domain- Blue, C-terminal domain ?Magenta, Large loop 1 (LL1) ? Orange, and 
Large  loop 2 (LL2) ? Green. 
 
37 
  
(98). Instead the space corresponding to heme pocket in this vestigial active site structure is filled 
with two small ? strands (between helices D? and E?) and a 10 amino-acid long loop (between 
helices F? and G?) (43, 98).  
Despite being ~30? away from the active site and inability to bind heme, the C-terminal 
domain is a crucial component in maintaining the bifunctionality of KatGs (86). Loss of the 
entire C-terminal domain results in complete loss of activity due to coordination of the active site 
histidine to the heme iron (86, 100). This observation was attributed to collapse of the actives site 
architecture due to disruption of interdomain interactions (86, 100). The same supporting 
function of C-terminal domain is performed by distal calcium ion in monofunctional 
peroxidases, such as lignin and manganese peroxidases (64, 80). One of the inter-domain 
interactions includes a strictly conserved hydrogen bond network formed between tyrosine from 
BC loop of the N-terminal domain, and the aspartate from B?C? loop of the C-terminal domain, 
has been shown to tune the KatG active-site for bifunctionality (101). Disruption of this 
hydrogen bond by substituting tyrosine with alanine yields an enzyme with diminished catalase 
activity (101). It is thus proposed that despite its distance from the active-site, the C-terminal 
domain serves to support an architectural framework necessary for proper active-site 
configuration by providing a series of interactions with N-terminal domain. 
In addition to the extra C-terminal domain, KatGs contain three large loops (LL1, LL2, 
and LL3) in the N-terminal domain. LL1 and LL2 are positioned at the opposite edges of the 
heme and restrict the access channel to the deeply buried active site at the distal side. LL3 is far 
from the active site and its role in KatG is not well defined (94).  
 
38 
  
LL1 spans between helices D and E and is located at one edge of the heme connecting its 
distal and proximal pockets (91-94). This structure varies considerably in length (36-76 amino 
acids) among KatGs, but it nevertheless contains a highly conserved sequence of ~10 amino acid 
residues ?MGLIYVNPEG? its C-terminal end (43, 102). Deletion of this loop results in elevated 
peroxidase activities compared to wild-type KatG with complete loss of catalase activity (103) . 
The strictly conserved residues of LL1, e.g., Tyr249, Ile248, and Glu253 (Synechocystis 
numbering), have been identified as essential for the H2O2 oxidation which is the hallmark of and 
unique to catalase turnover (91-94, 102). Glu253 creates an acidic entrance to the access channel 
which is characterized by a pronounced funnel shape and a series of water molecules (91-94). 
The substitution variants Glu253Gln sustains a substantial loss of catalase activity and alters the 
pH profile of catalase activity with no effect on peroxidase activity (43). This observation 
suggested that the negative charge of Glu253 at the entrance stabilizes and orients the solute 
matrix essential for H2O2 catalysis, particularly its oxidation (102, 104).  
The hydrogen bond between Ile248 of LL1 and Asp152 is important for the stability of 
the heme architecture, and perturbation of this hydrogen bond due to Ile248Phe mutation 
drastically reduces the catalase activity as a result of changes at the proximal heme site (105). 
These changes which are mediated by LL1 and its connection with helices E and F cause an 
increase in pentacoordinate high spin heme at the expense of hexacoordinate high spin heme 
form (105). Moreover, Asp152 has been shown to be essential for the catalase but not peroxidase 
activity of KatG (106). 
 
39 
  
All crystal structures for KatG solved so far reveal that the conserved tyrosine residue of 
LL1 is involved in a covalent adduct at the distal side of the heme. This adduct is formed 
between C?2 of tryptophan and C?1 of tyrosine, and between C?2 of tyrosine and S? of methionine 
(Figure1.14). The formation of this adduct is heme- and peroxide- dependent and is proposed to 
proceed by a radical-based mechanism initiated by the heme (107, 108). The formation of 
covalent adduct is also proposed to involve the tyrosyl and tryptophanyl radicals (109) but no 
direct insight about this has emerged to date. The existence of covalent adduct in solution has 
been shown by mass-spectrometry and SDS-PAGE (107, 110, 111). None of the three amino 
acids of the adduct can be substituted without diminishing the catalase activity of KatG, yet such 
mutants maintain or even exhibit improved peroxidase activity (92, 96, 108, 112). Thus, 
manipulation of this unique adduct affects the electronic nature of the redox intermediates. 
Mutation of tyrosine to phenylalanine has been shown to cause rapid conversion of compound I 
to stable compound II due to endogenous electron transfer (90, 113, 114). The formation of 
stable compound II favors the peroxidase activity but eliminates catalase activity (113, 115). The 
mutation also allows the easy conversion and stable formation of compound III, which interferes 
with the catalase activity (75, 90, 113, 114). Moreover, it has been observed that disruption of 
this adduct weakens the heme binding to the protein. This results in the appearance of 
hexacoordinate low-spin heme form, along with  the loss of the heme prosthetic group, due to 
binding of strong internal ligand to the heme iron (116). Recently, a scheme depicting the 
catalytic role of this adduct radical has been suggested in catalase reaction of KatG.  
 
 
40 
  
 
 
 
 
 
 
 
 
 
 
 
 Figure 1.14: Model of (Trp-Try-Met) covalent adduct of catalase-peroxidase 
 
 
 
 
H N
O H
S
?
R
R
R
M e t
C
? 2
C
? 1
C
? 2
T y r
T r p
 
41 
  
According to this scheme, tyrosine in the adduct can catalyze oxygen formation from 
superoxide, which dissociates from oxyenzyme during the catalase reaction of KatG (117).  
Other roles for LL1 and its relation to the covalent adduct will be addressed in Chapter Three. 
Sequence alignment performed using Clustal X (details presented in Chapter Four) on 20 
selected representatives from all known Class I families with one sequence of typical Class II 
representative shows that in contrast to LL1, LL2 varies little in its length (35-36 amino acids). 
However, in comparison to LL1, LL2 contains far fewer (~4 amino acids) invariant residues. 
This loop begins and terminates adjacent to the active site spanning between F and G helices, 
and its apex protrudes into a hydrophobic pocket in the C-terminal domain of the other subunit. 
A complete loss of catalase activity, 50% reduction in peroxidase activity, and diminished rate of 
cyanide binding was evident in the variant lacking the entire LL2 (87). However, the mechanism 
behind this observation was not clearly understood.        
Closer examination of the LL2 structure indicated that the ascending and descending 
limbs of this loop run roughly parallel to the hydrogen bond network formed by distal histidine 
(His106), Asn136, and Leu130. Thus, removal of entire LL2 was suggested to disrupt the above 
mentioned hydrogen bond network as two of its residues Glu284 and Trp297 lie fairly close to 
this network (87, 103). In order to determine the underlying mechanism of LL2 participation in 
the reactivity of KatG, the Goodwin lab has carried out a detailed study involving various amino 
acid residues of this loop. The data obtained from this study have been complied in Chapter Four 
of this dissertation. 
 
42 
  
The preceding discussion demonstrates that the large loops of KatG serve an important 
role in maintaining the catalytic versatility of the enzyme. To further evaluate these two large 
loops, they were subjected to modification by substitution and deletion mutagenesis followed by 
biophysical and kinetic characterization. For Large loop 1, the main goal was to investigate the 
additional functions of this large structure (36-76 residues long) besides positioning tyrosine 
correctly for the formation of the covalent adduct. In order to investigate this, various portions of 
the loop were deleted including or excluding tyrosine 226 and their catalytic properties were 
compared with wild-type KatG and its tyrosine to phenylalanine variants. 
In case of large loop 2, closer examination of this structure revealed an intersubunit 
interaction happening ~ 35? away from the active site between its apex and hydrophobic cavity 
of C-terminal domain of another subunit. Keeping in mind that the removal of entire LL2 
eliminated the catalase activity but preserved the peroxidase activity of KatG and also that C-
terminal domain is required to maintain both the activities of KatG, we focused on understanding 
the role of this intersubunit interaction in fine tuning the active site architecture and preserving 
the activities of KatG. LL2 was targeted in two different ways: first, important residues at the 
point of contact were either substituted or deleted, and secondly, the length of LL2 was targeted 
by deleting conserved and non-conserved residues. Both sets of mutations were made to disrupt 
the intersubunit interaction and to decipher the importance of length preservation in LL2 over 
residue preservation.  The detailed result and observation of this study is compiled in Chapter 
Four. 
 
 
43 
  
1.5. Conclusions 
Heme proteins are rich sources to exploit to understand the relationship between protein 
structure and function. The available three dimensional structures have provided crucial 
information about heme structure, location and role of important catalytic residues in and around 
the active site, effect of post-translational modification on protein?s catalytic ability and position 
and importance of peripheral structures of protein with respect to active site. The diverse 
functionality of heme proteins, even though they all contain small and simple metallo-organic 
compound heme, is fascinating and points to the factors which manipulate heme to achieve 
catalytic versatility. These factors include heme structure, protein-derived heme ligands, non-
ligand heme environment, and the global structure of the protein. Much interest has been paid to 
understand the effects of first three structural levels on heme protein activity, and the role of 
global structure of proteins has been under investigation. This thesis details my contribution to 
decipher the role of various peripheral structures and interaction arising from these structures on 
the active site environment and eventually on the activity of heme proteins. We have used 
Escherichia coli catalase-peroxidase (KatG) as a model to evaluate the mechanism by which 
structures peripheral to enzyme active site influence its architecture and catalytic ability. 
KatG catalyzes both catalase and peroxidase reactions using a unique active site. This 
active site catalyzes substantial catalase turnover even though it bears no resemblance to 
classical catalases. Instead, the active-site is superimposable on that of classical peroxidases, 
enzymes which have little if any catalase activity. This diversity in function highlights the effect 
of structures present outside the active site. Indeed, structural comparison of catalase-peroxidases 
 
44 
  
with classical peroxidases shows the obvious differences between two groups of enzymes like a 
gene-duplicated C-terminal domain and two interhelical insertions (LL1 and LL2) unique to 
catalase-peroxidases. This thesis focuses on understanding the role of these peripheral structures 
and interactions arising from them with the hope that it will shed some light on the regulation of 
heme protein function by global protein structure. 
 
 
45 
  
     Chapter Two: Generation and Characterization of Catalase-peroxidases by 
Molecular and Biophysical Techniques  
2.1. Manipulating the katG gene: cloning and mutagenesis 
The katG gene from E.coli (K-12) was cloned and placed in a pET expression vector by 
Dr. Douglas Goodwin using a PCR-based approach. Primers were designed to amplify the katG 
gene such that the product could be cleaved with Nde I at the 5? ATG start codon and by Xho I at 
the 3? end such that the natural stop codon was eliminated. The resulting PCR product and the 
pET-20b plasmid were each cleaved with Nde I and Xho I, and the correct fragments were 
ligated together according to standard procedure  (118).  The candidate plasmids were sequenced 
to eliminate the possibility of accidental mutations during cloning process.  By using the Xho I 
restriction enzyme with the pET-20b vector, the katG gene was followed by six codons for 
histidine and a stop codon.  Thus, wtKatG is expressed with a six-histidine tag on its C-terminus.   
The pET expression system is a system for the robust expression of recombinant proteins 
in E. coli. The pET expression vectors are designed to allow use of a T7 promoter for 
bacteriophage T7 RNA polymerase-based transcription of the target gene. This ensures that 
almost all of the cell?s resources are converted to target gene expression, but also minimize 
expression of the target gene until T7 RNA polymerase is itself expressed. In the BL-21(DE3) 
expression host, T7 RNA polymerase is under control of the lacUV5 promoter, and thus, its
 
46 
  
expression can be induced by the addition of IPTG.  
The pET vector used in the research described in this dissertation was pET-20b. The katG 
gene was inserted following the T7 RNA polymerase promoter. As mentioned above, the katG 
gene was inserted in such a way that the expression would result in a C-terminal six-histdine tag. 
For the selection purposes, the pET-20b plasmid encodes an ampicillin resistance marker, 
conferring ampicillin resistance on cells carrying the plasmid.  
 2.1.1. Site-directed mutagenesis 
In vitro site-directed mutagenesis is an invaluable technique for studying protein 
structure-function relationships and allows for the single substitution of amino acids purported to 
be critical to function.  
2.1.1.1. QuikchangeTM technique (Agilent Technologies) 
The QuikChangeTM approach (Agilent Technologies, La Jolla) was used to make point 
mutations that resulted in a change of the target codon. This method was performed using 
thermostable Pfu? DNA polymerase (Agilent Technologies). Pfu DNA polymerase was 
preferred as it replicates both plasmid strands with high fidelity and without displacing the 
mutant oligonucleotide primers (119). The Quikchange procedure utilizes a supercoiled double-
stranded DNA (dsDNA) vector (in this study recombinant His-tagged E.coli KatG (pKatG1) was 
used as a template) and two synthetic oligonucleotide primers containing the desired mutation 
(Figure 2.1). All oligonucleotide primers were purchased from Invitrogen (Carlsbad,CA) and  
 
47 
  
 
 
Figure 2.1: Overview of QuikChangeTM site-directed mutagenesis method. Scheme adapted 
from Stratagene QuikChangeTM site-directed mutagenesis manual. 
 
 
pka t G  1 i n pE T  20b w i t h t a rge t  s i t e  ( ) for m ut a t i on
P CR (D e na t ure s  t he   pl a s m i d a nd a nne a l  t he  pri m e rs  (     ) 
w i t h de s i re d m ut a t i ons
M ut a ge ni c  pri m e rs  w e re  e xt e nde d a nd i nc orpora t e d 
us i ng pfu D N A  pol ym e ra s e , t hi s  re s ul t e d i n ni c ke d c i rc ul a r
s t ra nds
M ut a t e d pl a s m i d w i t h ni c ke d c i rc ul a r s t ra nds . D pn I di ge s t i on 
at 37 
 
C for 1 - 2 hrs
T ra ns fe r t he  c i rc ul a r ni c ke d, ds  D N A  i nt o X L - 1 
Bl ue  s upe rc om pe t e nt  c e l l s
P a r e n t  D N A  P l a s m i d
M u t a g e n i c  P r i m e r
M u t a t e d  D N A  P l a s m i d
L e g e n d
 
48 
  
designed to be complementary to opposite strands of the vector, the mutation to be introduced 
and its complement were encoded to the middle of the coding strand and non-coding strand 
primers, respectively, and were extended during temperature cycling (PCR) by Pfu DNA 
polymerase. The resulting products contained staggered nicks, and these products were then 
treated with Dpn I endonuclease (New England Biolabs, Ipswich, MA) at 37 C for 1 hr. The 
target sequence (5?-G(Me)ATC-3?) of DpnI endonuclease is specific for methylated and E.coli 
hemimethylated DNA and is used to digest the parental DNA template and to select for DNA 
with desired mutation (120). DNA isolated from almost all E.coli strains is dam methylated and 
therefore susceptible to Dpn I digestion. The resulting plasmids were then used to transform the 
XL1-Blue non-expression host (Agilient Technologies) by electroporation using a Biorad 
MicropulserTM.  
2.1.1.2. ?Around the Horn? technique 
This is also a PCR-based method, but in contrast to QuikChange, it uses non-overlapping 
oligonucleotide primers to introduce mutations in the expression construct for the target protein 
(Figure 2.2). The primers are designed in such a way that if the 5? end of the forward primer is at 
the position ?x? in the sequence, the 5? end of the reverse primer is at position ?x-1?. The PCR 
products that result from the process have blunt ends and can be ligated using T4 DNA ligase 
(New England Biolabs) provided that primers are phosphorylated on their 5? ends. In all cases, 
ligation products were used to transform E.coli XL-1 Blue (a non-expression host) by heat shock. 
Candidate colonies were selected on the basis of ampicillin resistance. Plasmids from the 
candidate colonies were screened first by diagnostic restriction digests. Positive candidates were  
 
49 
  
 
 
 
Figure 2.2: Overview of ?Around the horn? site-directed mutagenesis method. 
 
 
 
 
.
.
pka t G  1 i n pE T  20b w i t h t a rge t  s i t e  ( ) for m ut a t i on
P CR (P l a s m i d de na t ura t i on a nd pri m e s  a nne a l i ng)
D pn I di ge s t i on a t  37 
 
C for 1 - 2 hrs
L i ga t i on of t he  m ut a t e d pl a s m i d 
 
50 
  
then submitted for DNA sequence analysis (Davis Sequencing, Davis, CA) to verify the correct 
mutations and ensure that no unintended mutations had occurred. Plasmids were used to 
transform E.coli BL-21 [DE3] pLysS (the expression host). 
 2.1.2. Deletion mutagenesis 
2.1.2.1. Seamless?cloning (Stratagene)  
  To generate the construct for the expression of deletion variants, a deletion protocol was 
developed based on Seamless?cloning (Stratagene).  This method is a PCR-based approach in 
which primers are designed to include Eam 1104 I restriction sites. Consequently all the products 
were treated with Eam 1104 I. Eam 1104 I is a type-IIS restriction endonuclease that cleaves 
target DNA downstream of its recognition sequence (121). The unique feature of Eam 1104 I is 
that it cleaves one nucleotide down on the upper strand in the 3? direction and four nucleotides 
down on the lower strand in 5? direction. This results in overhanging sequence with three 
nucleotides that can be designed into the primers to suit the investigators needs. 
 
5? ? CTC TTC N?NN  NN ? 3? 
3? ? GAG AAG NNN  N?N ? 5? 
  Primers were designed to amplify pKatG1 leaving out the codons coding for the 
targeted region, to include a Eam 1104 I restriction site, and yield compatible overhanging 
 
51 
  
sequences upon digestion with Eam 1104 I. The overhanging sequences were ligated using T4 
DNA ligase. The resulting products were used to transform E.coli XL-1 Blue cells by heat shock.  
Colonies were selected on the basis of ampicillin resistance, and candidate plasmids were 
verified by DNA sequence analysis at Davis Sequencing. 
2.1.2.2 ?Around the Horn? technique  
To generate deletion variants, non-overlapping primers were designed to exclude the 
codons corresponding to the targeted deletion. The desired codons were removed either from the 
forward primers or reverse primers. The desired PCR products were ligated using T4 DNA 
ligase. The resulting products were used to transform E.coli XL-1 Blue cells by heat shock. 
Colonies were selected on the basis of ampicillin resistance, and candidate plasmids were 
verified by DNA sequence analysis at Davis Sequencing. 
2.2. Expression system 
E.coli strain BL21-Gold[DE3]plysS was used to express the proteins studied in this 
dissertation. First the pET-20b plasmids containing the desired gene were used to transform 
E.coli strain BL21-Gold[DE3]plysS. The ?DE3? in the expression strain indicates that this strain 
is a lysogen of bactriophage DE3, implying that it contains a chromosomal copy of the gene for 
T7 RNA polymerase under the control of the lacUV5 promoter (122). Target genes are 
maintained in a transcriptionally silent mode until induction using the lactose analog, isopropyl-
?-D-thiogalactopyranoside (IPTG).  In the absence of IPTG, lac repressor binds to the lac 
 
52 
  
promoter, and little or no T7 RNA polymerase is expressed. Consequently, none of the T7 
promoter-controlled target protein is expressed. Conversely, in the presence of IPTG, the lac 
repressor dissociates from the lac promoter allowing for expression of T7 RNA polymerase and 
as a result the gene for the target protein is subsequently transcribed.  
The designation ?pLysS? indicates that a plasmid encoding T7 lysozyme is present in the 
strain. T7 lysozyme is a natural inhibitor of T7 RNA polymerase, and therefore, stops the 
?leakage? expression of the target protein (122). This way pLysS provides more tight control of 
target protein expression prior to induction. This is especially useful for the expression of toxic 
proteins. This provides additional stability to the target gene as even in the absence of IPTG, 
there is some expression of T7 RNA polymerase from the lacUV5 promoter in ?DE3 lysogen. If 
the target protein is sufficiently toxic to E..coli, the ?leakage? expression can substantially slow 
down the cell growth or cause cell death. As a result, the desired cell density for efficient and 
abundant expression of the target protein can be dramatically dimished. 
During transformation of the desired plasmid and expression host were incubated for 30 
minutes on ice. The mixture was then heat shocked at 42 C for 20 seconds and then placed on ice 
water for 2 minutes. Cells were grown in SOC medium for 1 hour with gentle agitation at 37 C. 
Resulting cells were plated on LB agar plates supplemented with ampicillin and 
chloramphenicol. Expression was carried out in 0.5 L Lauria-Broth with ampicilin and 
chloramphenicol at 37 C at 250 rpm. Expression was induced with IPTG (Gold Biotechnology 
Inc., St. Louis, MO) at mid-log phase (Optical density @ 600= 0.5) and cells were grown for 4 
 
53 
  
hours. At four hours post-induction cells were harvested by centrifugation at 10000 rpm at 4  C, 
supernatant was discarded and pellets were stored in -80 C for protein purification.  
Protein expression was analyzed by using the trichloroacetic acid (TCA) precipitation 
technique (123). A quantity of cells sufficient to yield a 0.05 OD600 reading (when diluted to 1 
ml) was treated with an equal volume of 10% trichloroacetic acid (4  C) followed by 
centrifugation. The pellet was then washed with 1 ml acetone, dried and resuspended in SDS-
PAGE loading buffer adjusting the pH with 200 mM Tris base. Protein solubility was monitored 
by taking 1 ml of cells from cultures at four hours post-induction of expression. Cells were 
centrifuged at 13,200 rpm for 10 minutes. Supernatant was discarded and pellet was 
homogenized with 75 ?l Bugbuster. The solution was centrifuged to separate out the pellet and 
supernatant. 10 ?l of supernatant was mixed with SDS loading buffer. Pellet was resuspended in 
40 ?l of urea, 10 ?l of urea suspended pellet was mixed with SDS loading buffer. Samples from 
expression and solubility analysis were separated by SDS- PAGE using 7.6% acrylamide 
resolving gel. 
2.3. Protein purification 
2.3.1. Soluble protein purification 
Cell pellets were resuspended in Bugbuster reagent (EMD4 Biosciences, Madison, WI) 
and 0.1ml phenylmethylsulfonylfluoride (PMSF). Bugbuster reagent is a protein extraction 
reagent and was used for the gentle disruption of the cell wall of E.coli to release soluble protein.  
PMSF is a serine protease inhibitor and was used to deactivate proteases from digesting target 
 
54 
  
protein after cell lysis. Following homogenization, 250 units of benzonase nuclease was added 
and mixture was incubated at room temperature for 2 hours with gentle stirring. Benzonase 
nuclease was used to remove nucleic acid contaminants from the protein preparation. This 
enzyme attacks and degrades all forms of DNA and RNA. The cell lysate was then centrifuged at 
10,000 rpm for 20 minutes. Since all the variants contain C-terminal six-histidine tag Nickel-
affinity chromatography was used to purify the protein. Nickel is bound to an agarose bead by 
chelation using nitrilotriacetic acid (NTA) beads. NTA is a tetradentate chelating adsorbent 
occupying four of six ligand binding sites in the coordination sphere of the nickel ion. The 
remaining two sites interact with six-histdine tag. Only the target protein containing six-histidine 
tag binds to resin as all other cellular proteins either do not bind or only weakly bind to the resin. 
The target protein is eluted with higher concentration of imidazole (Sigma, St. Louis, MO) 
whereas all other weakly bound proteins elute at lower concentrations of imidazole.   
 The supernatant obtained after the centrifugation of the cell lysate was loaded on to Ni-
NTA column (Qiagen,Valencia,CA) by circulating the solution at 1ml/min through the column 
bed overnight. Next day, the column was washed with 50mM Tris buffer, 50mM phosphate 
buffer followed by four different washes of 50mM phosphate, 200mM NaCl pH 7.0 buffer 
containing increasing imidazole concentrations ( 2mM, 20mM, 50mM, and 200mM). 2 and 
20mM washes removed most non-specifically bound proteins, whereas 50 and 200mM washes 
eluted the target protein off the column. Excess imidazole was removed by gel filtration 
chromatography using 50mM phosphate buffer and 50mM NaCl at pH 7.0. Protein was aliquoted 
in 1.5ml microcentrifuge tubes and stored at -80 C for future use.  
 
55 
  
2.3.2. Insoluble protein purification 
A denaturing protein purification procedure was used for proteins which were expressed 
in inclusion bodies. In this procedure, after cells were centrifuged down to pellets, the pellets 
were resuspended in 200 mM phosphate buffer, pH 7.0 (Buffer A) supplemented with 0.1 mM 
PMSF and 0.1 mg/ml lysozyme, and homogenized.  Cells were then lysed by sonication using 
Branson Sonifier 250 (Branson Ultrasonics Corp., Danbury, CT, USA).  Following sonication, 
benzonase (250 units) was added, and the mixtures were gently stirred at 23 ?C for 1 hour. The 
cell lysates were then centrifuged at 16,000 ? g.  The pellets, which contained the proteins of 
interest, were then resuspended in 8M urea and homogenized.  Centrifugation at 16,000 ? g was 
repeated. This time urea served to solublize the inclusion-body though they remained in a 
denatured state.  Thus, each supernatant was mixed with Ni-NTA resin and gently agitated at 23 
?C overnight.  The resin was then loaded into an empty column, washed with 10 mM imidazole 
in 8 M urea, and then the protein was eluted off the resin with 200 mM imidazole in 8 M urea. 
Urea and imidazole were then removed by dialysis against buffer A for 36 hours.  The bulk 
dialysis buffer was replaced every six hours. 
 2.4. Enzyme reconstitution 
After purification enzyme was reconstituted by addition of hemin (0.9 equivalents). Due 
to poor solubility of hemin at neutral and acidic solution, working solutions of hemin were 
prepared by dissolving solid hemin in 0.1 M KOH. After the addition of hemin, the enzyme was 
allowed to incubate for 24 hrs at 4 C. Following incubation, reconstituted enzyme was 
 
56 
  
centrifuged at 13,000 rpm for 5 minutes to remove insoluble hemin, complexes and protein 
aggregates. All spectral measurements for ferric and ferrous forms of enzyme were carried out 
by scanning the enzyme between 800 nm to 250 nm at room temperature in 100 mM phosphate 
buffer, pH 7.0 using Shmiadzu UV-1601 spectrophotometer. The pyridine hemochrome assay 
was used to determine the concentration of reconstituted enzyme (124). This assay is based on 
the characteristic absorption spectrum of oxidized and reduced form of the hemochrome by 
replacing the axial ligands with pyridine. Briefly, a reference solution of reconstituted enzyme, 
20% pyridine, 0.1 M NaOH and 0.3 mM K3Fe(CN)6 was made in a cuvette. A reference 
spectrum was then recorded from 560 nm to 530 nm. A miniscule quantity of sodium dithionite 
was then added and a second spectrum was recorded. Addition of pyridine, followed by the 
reducing agent (sodium dithionite) reduces the ferric enzyme to hexacordinate low spin ferrous 
form. Reconstituted enzyme concentration was calculated by the dividing the absorbance 
difference between the peak (~ 555 nm) and trough (~ 538 nm) absorbance by the difference in 
their absorptivities (20.7 mM-1 cm-1). 
2.5. Spectroscopic characterization of KatG variants 
  In order to understand the impact of substitutions and deletions within LL1 and LL2 on 
the structure and catalytic function of KatG, a full panel of spectroscopic and kinetic techniques 
was brought to bear on each one. Described here are the principles underlying each technique, 
the general approaches to our analyses of the data, and the information that each provides with 
regard to KatG structure and catalysis. The particular details of these experiments (e.g., 
 
57 
  
instrumental parameters, solution conditions and concentrations used, etc.) will be included in 
the subsequent chapters in which they are most appropriate. 
2.5.1. Circular dichroism (CD) spectroscopy 
 Circular dichroism occurs when a sample differentially absorbs left-handed circularly 
polarized (lcp) light and right-handed circularly polarized (rcp) light (??=?L-?R). This is a 
property of a chiral or optically active chromophore. Most biological macromolecules, including 
proteins, are composed of chiral building blocks and these components often have chromophores 
in the UV range. CD spectroscopy is a technique in which the CD of a molecule is measured 
over a range of wavelengths. For proteins, the most frequent application of this technique is to 
evaluate CD at far UV wavelengths to assess secondary structural content (e.g., ?-helix, ?-sheets, 
etc.). Individually, the n??? and ???? transitions associated with the peptide bond are very weak 
and tend to cancel one another. However, in a given secondary structure where the 
conformational features of peptide bonds are repeated, a cumulative effect is observed which is 
characteristic for the major types of secondary structures found in proteins. 
Effective application of CD spectroscopy depends on comparison against some known 
standard. In case of using CD as a measure of protein structural stability (with respect to 
temperature, pH, or otherwise), spectra recorded under conditions favoring the unfolding of a 
protein are compared against the spectrum recorded under conditions favoring the folded state. 
Likewise, a CD spectrum for wild-type protein is the standard against which any variant is 
 
58 
  
measured. Finally, the secondary structural content of an unknown protein can be estimated from 
its spectral characteristics when compared against CD spectra for proteins of known structure. 
Secondary structure of the proteins is determined in the ?far-UV? spectral region (190-
250 nm). Different secondary structures of proteins such as ? helices, ? sheets, random coil, type 
I ?-turn, and extended 310-? helices give rise to characteristics shapes and magnitude of CD 
spectrum (Figure 2.3) (125).  The ?near-UV? spectral region (250-350 nm) CD spectrum of a 
protein is helpful for determining certain features of tertiary structure. Aromatic amino-acids and 
disulphide bonds at this wavelength range produce CD signals which are sensitive to the changes 
in the overall tertiary structures of  proteins. In this study, all the variants prior to reconstitution 
with hemin were used to record the far-UV CD spectra (180 -300 nm) using a Jasco J-180 
spectrapolarimeter. To ensure that absorption by samples was not excessive, low concentrations 
(5 ?M) of protein were prepared in only 5mM phosphate buffer and a minimal pathlength (0.05 
cm) CD cuvette was used. Phosphate buffer was used as it is one of the better buffers in terms of 
its minimal absorption in the UV. Low concentration of sample and buffer ensured maximal 
signal to noise ratio in the CD spectrum and avoided masking of CD signals by the buffer. The 
short pathlength of cuvette helped to overcome sample absorption and permitted scanning to 
lower wavelengths. The instrument was purged with N2 gas in order to remove O2 from the lamp 
housing and the sample compartment which prevented ozone formation and minimized the 
damage to the optical systems. Also, N2 atmosphere allowed measurements to be made below 
200 nm. In order to improve the quality of the data and to be consistent in data acquisition over 
the range of variants, experimental parameters like the time constant, the scan rate, the number of 
scans, and the bandwidth were kept standard.  
 
59 
  
 
 
Figure 2.3: Far UV CD spectra associated with various types of secondary structures. Solid 
line, ?-helix; long dashed line, anti-parallel ?-sheet; dotted line, type 1 ?-turn; cross dashed line, 
extended 310- ?-helix; short dashed line, irregular structure (125). 
 
60 
  
2.5.2. UV-visible absorption spectroscopy 
 Although UV-visible absorption is central to a number of our kinetic studies, the kinetic 
application of UV-vis will be covered later in this chapter. As a purely, spectroscopic tool, UV-
visible absorption has two primary applications. The first is to evaluate KatG as a generic protein 
based on absorption due to the presence of aromatic amino acid side chains, particularly 
tryptophan and tyrosine. Phenylalanine does also absorb UV light, but due to its very low molar 
absorptivity, its contribution to the overall spectrum of KatG is minimal. Likewise, cysteine is 
recognized to be a contributor, but E.coli KatG has only one cysteine side chain. The dominant 
band in the UV has an absorption maximum at 281 nm. This is the result of a cumulative 
contribution of 22 tryptophan residues and 15 tyrosine residues, resulting in a particularly and 
unusually intense absorption band. Indeed, the theoretical molar absorptivity expected from this 
composition is 144,000 M-1cm-1 (126). The intensity of absorption at 280 nm is a convenient 
method of estimating the concentration of purified KatG. The spectrum for KatG from 320-250 
nm also serves for the qualitative evaluation of the stability of the protein isolated. A 
progressively intensifying baseline as one scans further into the UV is indicative of light 
scattering. Though there are several underlying causes for light scattering in a given sample, one 
particularly relevant to the work described here is protein aggregation. Substantial light 
scattering suggestive of protein aggregation is an early indication that a given variant lacks 
structural stability and is prone to non-specific association and precipitation. 
  Because KatG is a heme-containing enzyme, UV-visible absorption is also highly useful 
for evaluating the oxidation state and coordination environment of the heme itself. Generally, 
 
61 
  
speaking, heme will have at least three prominent absorption bands which arise due to ?-?? 
transitions in the conjugated ? system of the heme. In order of increasing energy and intensity, 
these bands are called ? (Q00), ? (Q0v), and ? (Soret). The Soret band (?) is the most intense band, 
and is the highest energy transition therefore, occurs at lowest wavelength of the three. In free 
heme, the Soret band is observed at 380 nm. When incorporated in a protein, the absorption will 
red shift typically to wavelength greater than 400 nm. When electrons from the porphyrin ?-
system are transferred from highest energy-filled orbital to lowest energy unfilled orbitals (d?) of 
the iron, charge-transfer (CT) bands are observed. The appearance of CT bands depends upon the 
spin state of heme.  The characteristic absorption bands of KatG in its resting state occur at 408 
nm (Soret), 502 nm (CT2), and 640 nm (CT1) (Figure 2.4).  In the high-spin complex of KatG 
the ? and ? bands are largely obscured by the charge transfer bands. However, charge transfer 
bands are not observed in low-spin complexes and are too weak to observe when the iron is in 
ferrous state. In low-spin complexes, the charge transfer is blocked due to dropping of a 
porphyrin ?? electron into a d orbital of the iron completely filling the d orbitals. The 
characteristic absorption bands of the low-spin ferricyano-KatG complex occur at 422 nm 
(Soret), ~542 nm (?), and ~575 nm (?). Thus, it is evident that the ligand environment of heme 
iron affects its optical absorptions, both at ?-?? and CT transitions. 
Isolated KatGs contain heme with the iron in its ferric oxidation state. Ferric iron 
contains five d-electrons, in the absence of external influences and according to Hund?s rule, 
each occupy one of five degenerate d-orbitals (Figure 2.5). Though a somewhat rudimentary 
approach, crystal field theory provides a straightforward explanation for the effect of ligands on  
 
62 
  
325 350 375 400 425 450 475 500
0
15
30
45
60
75
90
105
H i g h - s p i n  F e r r i c
F e r r i - Cy a n o
L o w - s p i n  F e r r i c
W a v e le n g t h  ( n m )
?
 
(
m
M
-1
cm
-1
)
500 525 550 575 600 625 650 675 700
0 . 0
2 . 5
5 . 0
7 . 5
1 0 . 0
1 2 . 5
1 5 . 0
Hi g h - s p i n  F e r i c
F e r r i - Cy a n o
L o w - s p i n  F e r r i c
W a v e le ng t h ( nm )
?
 
(
m
M
-1
cm
-1
)
 
Figure 2.4: Typical absorption spectra for KatG in the Soret (top) and charge transfer 
(lower) regions. 
 
63 
  
the energies of the five d orbitals. Each of six ligands can provide a lone pair of electrons for 
bonding to the iron, and in an octahedral complex could occur along the x, y, and z axes. By 
virtue of electron-electron repulsion, iron d orbitals oriented along the axes (i.e., dx2-y2 and dz2) 
will be higher in energy than those oriented between major axes (i.e., dxy, dxz, and dyz). Ligands 
which destabilize the on-axis orbitals in such a manner that electron pairing energy is less than 
the energy required to place an electron in a destabilized orbitals (i.e, splitting energy) are strong 
field ligands (Figure 2.5) whereas, the ligands which do not split the energy levels so severely 
are weak field ligands. In the presence of strong field ligand iron (III) has a +? spin (i.e., low 
spin) whereas in the presence of weak-field ligand it will have +5/2 spin (i.e., high spin).  
Other factors can result in further differentiation of the d orbital energies, and such is the 
case in many heme proteins (Figure 2.6). The pyrrole nitrogens of heme are located in the plane 
of dx2-y2 and dxy orbitals of iron and electrons in dx2-y2 are pointed towards the bonding pair of 
electrons of pyrrole nitrogens and are strongly repelled. Also, the electrons in dxz and dyz orbitals 
are repelled more strongly than in dxy due to the conjugation of the porphyrin. The dz2 orbital is 
sensitive to the fifth and sixth ligands, both of which are located along the z-axis. As stated 
earlier, in the conjugated ? system of the heme, three ?-?? transitions designated as ?, ?, and ? are 
observed. The ? transition originates from a2u state to ?? (eg) state.  The ? band has the same 
origin but goes to the vibrationally excited levels of the eg state, and thus is of slightly higher 
energy. The Soret band (?) is of the highest energy due to the fact that it originates from the 
lower energy ? (a1u) ground state but ends at the same eg ?? as the ? transition (Figure 2.7). 
 
 
64 
  
 
 
 
Figure 2.5: Effects of ligands on FeIII d-electrons. 
 
 
 
No  field W e a k  li g a n d  field S tr o n g  li g a n d  field
 
65 
  
 
 
d
xy
d
xz
, d
yz
d
z
2
d
x
2 
-
y
2
 
 
Figure 2.6: Further splitting of d-orbital energy levels in heme peroxidases. 
 
 
66 
  
 
 
 
 
Figure 2.7: Electronic transitions observed in heme peroxidases.  
 
b
2g
(d ? ) e
g
a
1g
b
1g
B (S o re t )Q
0v
( ? )
e
g
( ? ?)
a
2u 
( ? )
a
1u 
( ? )
Q
00
( ? )
CT
I r on H e me
En e r gy
 
67 
  
2.5.3. Magnetic Circular Dichroism spectroscopy 
Magnetic circular dichroism (MCD) as the name suggests is the measurement of a CD 
signal in the presence of external magnetic field oriented parallel to the direction of light 
propagation. The magnetic field or Faraday effect is induced in a particular chromophore through 
its effect on the electronic properties of the absorbing species as a consequence of Zeeman 
splitting. Three different types of MCD spectra are evident and are termed as A, B, and C. A and 
B terms are temperature-independent, whereas the C term is temperature dependent (127, 128).  
An ?A term? arises when transitions occur from a non-degenerate ground state to a 
doubly degenerate excited state (Figure 2.8). These transitions require absorption of lcp and rcp 
light giving rise to derivative shaped signal. A ?C term? is same as an ?A term? but instead of 
magnetic field splitting, it splits the ground state (Figure 2.9). This causes difference in the 
intensity between lcp and rcp resulting in Gaussian shaped signal. A ?B term? arises from 
nondegenerate ground-state and excited-state mixing brought on by the magnetic field and is 
Gaussian in shape. 
MCD spectra have been observed for numerous heme proteins of known oxidation and 
spin state. This provides a substantial archive for comparison to new and lesser known heme 
systems like those of KatG. Moreover, the identity of axial ligands and characteristics of their 
protein environment can be evaluated, but this typically requires access to near infrared 
wavelengths. The easiest application of MCD to heme proteins is distinguishing between high-
spin and low-spin ferrous heme which can be accomplished by monitoring features between 350 
and 750 nm. 
 
68 
  
 
 
 
Figure 2.8: Basis for an MCD spectrum: A-term. 
 
 
1
P
+1
l .c .pr .c .p
1
P
- 1
1
P
0
1
P
1
S
1
S
0
A
b
s
or
b
an
c
e
?
?
=
?
l
-
?
r
-
+
A - te r m
 
69 
  
 
 
Figure 2.9: Basis for an MCD spectrum: C-term. 
 
 
1
P
+1
1
P
- 1
1
P
0
1
P
l .c .pr .c .p
1
S
1
S
0
A
b
s
or
b
an
c
e
?
?
=
?
l
-
?
r
-
+
C - te r m
 
70 
  
The transitions observed in the MCD of ferrous heme are due to the Soret band and ? and 
? bands. An intense ?A? term signal is observed near 550 nm in the low-spin ferrous heme due to 
the ? and ? band. However, in case of high-spin ferrous heme and also low and high spin ferric 
heme, the spectra are more complex. This is due to the presence of new transitions like charge 
transfer bands (and possibly weaker d-d transitions) and the unpaired electrons on the iron which 
produce a spin degeneracy in the system. Two effects arise due to this ground state spin 
degeneracy; first a C-term is produced, and second, spin-orbital coupling of iron to porphyrin is 
observed, which splits the porphyrin energy level (129, 130). According to spin-orbital coupling 
theory high-spin ferrous heme MCD spectrum show Soret absorption maxima near 430-440 nm 
and exhibit an intense positive MCD and weaker negative MCD signals. Also the features near 
550 nm are less intense in case of high-spin ferrous heme. The difference in the intensity due to 
the ? and ? band near 550 nm and near absence of Soret maxima in case of low-spin ferrous 
heme around 435 nm is a good marker for distinguishing the contribution of high- and low-spin 
ferrous heme (Figure 2.10). 
2.5.4. Electron Paramagnetic Resonance (EPR) 
Optical spectroscopy is a highly effective and straight forward method for the 
determination of concentration and enzymatic activity. However, its use as a tool for structural 
characterization of metal sites in proteins is far more limited. In particular, for heme proteins it 
should be remembered that the majority of spectral features observed by UV-Vis are due ? to ?? 
transitions in the porphyrin ? electron system rather than the metal itself. EPR spectroscopy 
circumvents these issues and is an ideal technique to characterize the structure and function of  
 
71 
  
 
 
 
370 470 570 670
- 1 0 0
- 5 0
0
50
100
439
550
W a v e leng t h  ( n m )
??
/
H
 (
M
-1
cm
-1
T
-1
)
 
 
 
Figure 2.10: Characteristic differences in ferrous MCD spectra of high-spin (black) and 
low-spin heme (gray). 
 
 
 
 
72 
  
heme proteins. In heme proteins EPR usually applies to their ferric form and in this state two 
characteristic families of EPR spectra predominate; low-spin Fe3+ (S=1/2) and high-spin Fe3+ 
(S=5/2) (131). 
The basic principle behind EPR relies on the interaction of a paramagnetic species within 
the molecule of interest with an external magnetic field. The external magnetic field lifts the 
degeneracy of the + ? and ? ? electron spin states such that one is aligned with the magnetic 
field (ms = + ?) and the other is aligned against the magnetic field (ms = - ?) (Figure 2.11). The 
energy difference between the two spin states is given by the equation 2.1, which is dependent 
upon the strength of the magnetic field (B) (132). 
? E = E+ - E- = h? = g?B     (2.1) 
Where B is the strength of the magnetic field in Tesla but usually represented as Gauss, ? is the 
Bohr magnetron (9.274 x 10-24 J T-1), h is Planck?s constant (6.626 x 10-34 J s-1), and ? is 
operating frequency of the spectrometer radiation. Of course, h and ? are constants and in the 
EPR experiment, and the frequency of radiation is held constant at a given level depending on 
the band EPR (e.g., ? at X-band is 9 GHz). The g-factor is the Zeeman splitting factor, and it is a 
tensor which specifies the strength of interaction between unpaired electrons and the applied 
magnetic field, B. In a paramagnetic species, the g-factor depends on the coordination number 
and geometry of the paramagnetic center and the chemistry of the primary ligands to the heme. 
This indirectly provides information about the location and environment of the unpaired 
electrons.  
In paramagnetic species, the g-factor is anisotropic meaning that it depends directly on its 
 
73 
  
 
Figure 2.11: Electronic paramagnetic absorption. 
 
M
s
+ 1/ 2
- 1/ 2
En e r gy (E)
B= 0 B
?
?
E
?
=  + g ? B/ 2
E
?
= - g ? B/ 2
? E  =  h v
A b s or p ti on
1
st
D e r i vati ve
 
74 
  
orientation in the magnetic field. Thus, for anisotropic species, the g-factor breaks down into 
different values of g for different orientations depending on the alignment of B along the 
Cartesian axes (x, y, and z) (39). For instance, in case of a heme protein, if the magnetic field is 
perpendicular to the plane of heme porphyrin it can result in three g-values gx, gy, and gz, and 
there are three types of symmetry associated with these three g-values (Figure 2.12) (131, 133). 
When all three g-values are equal (gx = gy = gz), the symmetry around heme is isotropic and is 
not dependent on orientation in the magnetic field. When two g-values are equal, the resulting 
EPR spectrum is axially symmetric (suggesting an axially symmetric microgeometry for the 
paramagnet). The unique g-values are g? (parallel to the symmetric axis =gz) and the other two 
equivalent g-values are referred to as g?. The axial symmetry has two subsets. One where g? > g? 
and second where g? > g?. When all the three g-values are unequal (gx ? gy ? gz) it refers to 
rhombic case and implies that the microsymmetry around heme iron is no higher than rhombic. 
In KatG three EPR spectra are generally observed; hexacoordinate high-spin ferric heme 
spectrum is dominated by axial with slight contribution from rhombic signal (Figure 2.13, Panel 
A), pentacoordinate high-spin ferric heme dominated by rhombic signal with minor contribution 
from axial signal (Figure 2.13, Panel B) and hexacoordinate low-spin ferric heme spectrum 
(Figure 2.13, Panel C). The EPR of high-spin ferric heme have g-values centered around g?=6 
and g? =2, and in low-spin EPR ferric heme three different values of g are seen (131). 
For the purpose of the work here, EPR was used as a tool to determine the effect of 
various mutations on spin state and coordination environment of the heme in the active site. It 
was also used to determine the relative concentrations of the different species in the population. 
 
75 
  
 
 
 
 
 
Figure 2.12: EPR absorption derivatives. 
 
 
 
 
76 
  
1000 2000 3000 4000
6 . 0 0
1 . 9 9
4x
A
M a g n e t ic F ield  ( G )
1000 2000 3000 4000
6 . 6 3
4 . 9 5
1 . 9 5
4x
B
M a g n e t ic F ield  ( G )
1500 2500 3500 4500 5500
2 . 9 2
2 . 2 8
1 . 5 3
C
M a g ne t i c  F i e l d ( G )
 
Figure 2.13: Different EPR spectra observed in catalase-peroxidases. High-spin 
hexacoordinate heme (A), high-spin pentacoordinate heme (B), and low-spin hexacoordinate 
heme (C). g-values shown in each spectrum are the estimates and can vary to some extent 
depending upon the heme environment. 
 
77 
  
To obtain sufficiently intense signals for each EPR spectrum, frozen samples of ~300 ?L 
at ~150?M concentration were prepared in liquid nitrogen and spectra were recorded at low 
temperature (~10 K). EPR signal magnitude is proportional to population difference, which is 
dictated by a Boltzman distribution. Low temperature increases the population difference. But 
this population difference is depleted by microwave excitation and restored by intrinsic spin-
lattice relaxation process. Thus, low temperature increases the EPR amplitude and decreases the 
linewidth by decreasing the spin-lattice relaxation rate, which in turn increases the lifetime of the 
excited state, and decreases the uncertainty in the excited state (132). 
The double integration of the absorption derivatives of the individual components was 
used to quantify the relative amount of each species in EPR spectra.  The resultant spin intensity 
was compared with that of a standard to calculate concentrations of species present. In order to 
determine how the intensities of the EPR signals obtained from KatG variants were related to the 
concentrations of the paramagnetic species responsible, spin quantification was carried out using 
Biomolecular EPR spectroscopy software package available online (134). The strategy was to 
obtain the pure line shapes of all the signals, integrate these line shapes, and then finally measure 
how much of each there was in every spectrum.  
In KatGs, EPR signals were mainly characterized by three effective g-values; two for 
high-spin heme species (axial and rhombic) and the third one for low-spin heme species. In some 
cases, axial signal showed two subsets with g-values very close to each other which complicated 
the spectrum and hence quantitation of different species. Thus, three high spin signals and one 
low-spin signal were required to obtain the desired simulations.  The high spin signals included: 
two axial signals (AHS1) with g-values of 5.85 and 1.99, and AHS2 with g-values of 5.69 and 
 
78 
  
1.99 and one rhombic signal (RHS) with g-values of 6.65, 4.95, and 1.95. The low-spin signal 
was rhombic (RLS) with g-values of 2.93, 2.27, and 1.53. After obtaining the lineshapes of all 
the individual signals, double integration was carried out. From these integration values, 
percentages of each paramagnetic species were calculated.  
2.6. Kinetic characterization of KatG variants 
The spectroscopic tools reviewed above provide valuable insight into the environment 
around a protein bound heme and the effect of amino acid substitutions or deletions. However, to 
be of interpretive value these effects must be related to the catalytic performance of the enzyme 
in question. Steady-state kinetic methods allow for the evaluation of catalytic performance under 
multiple turnover conditions monitoring the continuous generation of product or consumption of 
substrate. These studies allow for the determination of general parameters governing enzyme 
turnover as a function of substrate concentration. Transient-state kinetic methods allow one to 
evaluate individual steps in the catalytic process to determine the basis for changed steady-state 
kinetic parameters. 
2.6.1. Steady-state kinetic analysis 
Catalase and peroxidase activities 
One catalase activity assay measures the decomposition of H2O2 to H2O and O2 in the 
presence of enzyme at 240 nm (?(H2O2) at 240 = 39.4 M-1 cm-1) (135). This assay is typically 
carried out at room temperature using pH 7.0 phosphate buffer. 
 
79 
  
For the peroxidase activity assay, H2O2 is reduced and suitable electron donor is 
subsequently oxidized. The oxidized electron donor absorbs light and can be determined 
spectrophotometrically. Because the identity of the physiological electron donor of KatG is still 
unknown, a wide variety of organic and inorganic substances serve as electron donors. In this 
work, ABTS (2,2?-azino-bis[3-ehtyl benzothiazoline-6-sulphonic acid]) was used as the electron 
donor. ABTS was chosen because its radical oxidation product relatively stable in aqueous 
solution, has a high molar absorptivity at 340 nm (?340 = 3.66 x 104 M-1 cm-1) and its oxidized 
form absorbs at 417 nm (?417 = 34.7 mM-1 cm-1) (136). The data from the activity assays were fit 
to one of the following equations.  
When no substrate-dependent inhibition was observed, equation 2.2 (i.e., Michaelis-
Menten equation) was used (137).    
             
    
In this equation, v is initial velocity of formation of product, [E]T  is the total enzyme 
concentration, [S] is substrate concentration, kcat (turnover number) is the maximum number of 
substrate molecules converted to products per active site per unit time, and KM (the Michaelis 
constant) is the concentration required to reach half the maximum velocity. At saturating 
concentrations of substrate v/[E]T is equal to kcat, whereas at low substrate concentrations, the 
slope of v/[E]T vs [S] approaches the apparent second order rate constant kcat/KM. kcat/KM  is a 
measure of efficiency of the enzyme with a given substrate. 
v = k c a t  [ S ]
K M  +  [ S ][ E ] T
( 2 . 2 )
 
80 
  
This classical Michaelis-Menten equation describes the simplest enzymatic mechanism in 
which the association of enzyme with substrate is followed by the release of product (equation 
2.3) (137): 
      
 E - free enzyme, S ? substrate, ES ? the enzyme-substrate complex, P ? product, (k2 + k-1) / k1 = 
KM, and k2 =kcat 
The mechanism of KatGs is quite complex such that the standard interpretation of 
Michaelis-Menten parameters cannot be applied even though the catalase activity of KatG does 
follow the Michaelis-Menten kinetics allowing reasonable estimation of apparent kcat and KM 
values. However, peroxidase kinetics pose an additional difficulty due to substrate-dependent 
inhibition with respect to both H2O2 and some electron donors. Thus, in this situation it becomes 
difficult to observe true saturating conditions. In this case, equation 2.2 was modified to include 
substrate-dependent inhibition to accommodate a second apparent dissociation constant (KI) 
corresponding to the substrate?s action as an inhibitor. Hence, equation  2.4 was used when 
substrate dependent inhibition was detected (137). 
 
 
In its simplest form this relationship is seen when a second molecule of substrate binds to 
ES to give an ES2 complex that is catalytically inactive, equation 2.5 (137): 
E  +  S E S E  +  Pk 1k - 1 k 2 ( 2 . 3 )
=
k c a t  [ S ]
K M  +  [ S ]  +  [ S ] 2   K I
( 2 . 4 )v
[ E ] T
 
81 
  
 
 
 
 
At low concentrations of substrate, rates can still be fit to a Michaelis - Menten equation. 
But as [S] increases, there is first a maximum value of v followed by a decrease.  
2.6.2. Transient-state kinetic analysis of KatG 
 Stopped-flow experiments 
Steady-state kinetic measurement usually defines the kinetic boundaries of an enzyme-
catalyzed reaction by providing two essential pieces of data on kinetic performance, one 
corresponding to low concentration of substrate (kcat/KM) and one corresponding to high 
concentration of substrate (kcat). In order to measure the rate constants of the individual steps on 
the reaction pathway and to detect and monitor intermediates involved in the catalytic process, it 
becomes necessary to measure rates by transient-state methods.  
Transient-state kinetic methods are usually applied to the observation of rates that occur 
in a very short time (i.e., the short period of time from the enzyme encountering the substrate / 
inhibitor / other ligands to the point of equilibrium). Our instrument can be used to detect and 
monitor intermediates occurring in this time frame. Stopped-flow apparatus can be used to carry 
out either single mixing or double mixing experiments. In the work described in this dissertation, 
stopped-flow was used to observe the reactions of H2O2 and ABTS with large loop 1 variants and 
E  +  S E S E  +  P
k 1
k - 1
k 2
+
S
E S 2
K I
( 2 . 5 )
 
82 
  
also to monitor the rate of cyanide binding in case of large loop 2 variants. A SX.18MV rapid 
reaction analyzer (Applied Photophysics Surrey, UK) was used to carry out all the stopped-flow 
measurements. The basic set up of the instrument is shown in Figure 2.14 (138).  
In single-mixing experiment, the drive motor rapidly fires two solutions (e.g., enzyme in 
syringe A and substrate in syringe B) together into a mixer. The mixed solution then flows into 
the observation cell. At this point, a stop syringe limits the volume of the solution used in each 
experiment and stops the flow of the solution. The flow of the solution into the stop syringe 
moves the plunger back and triggers the data collection. The mixed solution in the observation 
cell is illuminated by a light source and the change, as a function of time, in absorbance can be 
measured.  
For large loop 1 variants in single mixing experiments, the enzyme was allowed to react 
with varying concentrations of H2O2 for a defined period of time. This allowed for the formation 
of higher oxidation states which were monitored initially by diode array to observe the formation 
and transition of different oxidation states from the native ferric state. Later, rate of formation of 
these different states were recorded at relevant single wavelengths. The single wavelength was 
chosen carefully to get an optimal distinction between formation and decay of one species to 
another. 
For large loop 2 variants, the enzyme was allowed to react with varying concentrations of 
KCN. Full spectra were recorded by diode array to observe the shift in the Soret band as enzyme 
with high-spin ferric heme species transitioned to low-spin ferric heme form (Figure 2.15). 
The kinetic constants for cyanide binding were measured by observing the time course of 
 
83 
  
Single-Mixing Stopped-Flow 
F l u or e s c e n c e
O p ti c al  P ath A b s or b an c e
M i xe r
D r i ve
S top p i n g
S yr i n ge
A B
C e l l
 
Double-Mixing Stopped-Flow 
F l u or e s c e n c e
O p ti c al  P ath A b s or b an c e
M i xe r  1
D r i ve  1
S top p i n g
S yr i n ge
A B
C e l l
D r i ve  2
C D
M i xe r  2
D e l ay Li n e
 
Figure 2.14: Basic setup for stopped-flow instrument. Scheme adapted from 
http://www.hi-techsci.com 
 
84 
  
cyanide binding at single wavelength (either 400 or 428 nm). Observed pseudo-first order rates 
were plotted as a function of cyanide concentration to obtain second-order rate constants for 
cyanide binding.  
In double mixing experiments, three reactants are mixed together in a sequential fashion. 
The first two syringes (enzyme in syringe A and one substrate in syringe B) drives the two 
solutions together through mixer 1 into a delay line. The time between the first and second ram 
firing determines the length of the first reaction. After passing through the delay line, the 
solution enters the mixer 2 where the third reactant is encountered. Then the mixed solution is 
passed into the observation cell. This triggers the data collection as described for the single 
mixing experiment. 
In double mixing experiments carried out using large loop 1 variants, enzyme was first 
allowed to react with H2O2 and the reaction mixture was left to incubate in delay line for set 
period of time. The content of delay line was then mixed with ascorbate and varying 
concentration of ABTS. The full spectra were recorded by diode array and rate constants for 
ABTS-dependent reactions were measured at single wavelength. Ascorbate was used as it 
reduces ABTS radicals as rapidly as they are generated, ensuring a constant substrate 
concentration during the reaction (114, 139). Ascorbate is often a poor substrate for peroxidase 
enzymes and has been a very effective tool for measuring rapid peroxidase reactions with 
electron donors and minimizes the interference from the oxidized products. Moreover, the 
concentration of ascorbate was kept constant in our ABTS stopped-flow studies making its 
contribution easy to account for. 
 
85 
  
 
 
 
325 350 375 400 425 450 475 500
0 . 0 0
0 . 0 3
0 . 0 6
0 . 0 9
0 . 1 2
0 . 1 5
0 . 1 8
0 . 2 1
408
422
413
A
W a v e l e ng t h ( nm )
A
bs
o
r
ba
nc
e
475 500 525 550 575 600 625 650 675 700
0 . 0 0 0
0 . 0 0 5
0 . 0 1 0
0 . 0 1 5
0 . 0 2 0
0 . 0 2 5
0 . 0 3 0
0 . 0 3 5
540
642
B
W a v e l e ng t h ( nm )
A
bs
o
r
ba
nc
e
0 5 10 15 20 25 30
0 .1 0
0 .1 2
0 .1 4
0 .1 6
0 .1 8
0 .2 0
A b s
400
A b s
428
C
T im e  ( s )
A
bs
o
r
ba
nc
e
 
Figure 2.15: Spectral changes (A and B) and typical time traces (C) observed for reaction 
of ferric enzyme with potassium cyanide. The kinetic constants can be measured by observing 
the decrease in absorbance at 400 nm or increase in absorbance at 428 nm, as both wavelengths 
fall on either side of the isosbestic point (413 nm). 
 
86 
  
Rapid Freeze Quench (RFQ) EPR experiment 
In principle, EPR spectroscopy can be used as a method to monitor kinetic event; 
however, in practice, the time required to tune the spectrometer, and its intrinsically low 
sensitivity compared to light-absorption spectroscopy, affects its competitiveness. This problem 
has been overcome by using rapid mixing-freezing method to study the reaction intermediates. 
This method of quenching a chemical or enzymatic reaction by rapid freezing was first 
introduced by Bray in 1961 (140) and adapted specifically for EPR analyses of the samples 
(141). RFQ-EPR can provide electronic and molecular structural information on biochemical 
transient intermediates with high time resolution, formed during the reaction whose presence was 
determined first by stopped-flow experiments. By preparing a series of freeze-quenched samples 
at various time intervals, a full kinetic profile of the catalytic cycle of an enzyme can be 
obtained.  
The freeze quench experiments are divided into three phases: the mixing, the ageing and 
finally the freezing (132). The frozen reaction products are collected and analyzed by EPR. 
Briefly, the procedure consists of the rapid mixing of two reactants (e.g., enzyme and substrate), 
followed by rapid freezing of the mixture in a liquid isopentane bath. The reaction is aged for a 
user-defined time inside the calibrated ageing loops. A built-in ejection nozzle at the exit of the 
ageing loop sprays the aged solution towards the dewar, which is filled with isopentane. The 
temperature is regulated by immersing the dewar in a liquid N2 bath. Cryosolvent level is 
maintained as close as possible to the nozzle to minimize flying time. On ejection the reaction is 
stopped below -130?C and the frozen sample is packed directly into the EPR tubes and analyzed  
 
87 
  
by EPR spectroscopy. 
In KatG, compound I is formed upon reaction of ferric enzyme with alkyl peroxides 
(142). The presence of this compound I species have been observed in various KatGs using 
optical stopped-flow experiment (75, 104, 113, 143, 144). This compound I decays quickly to 
compound II like species in the absence of exogenous substrates. This is consistent with an 
endogenous electron transfer pathway producing a tyrosyl radical on tyrosine involved in W-Y-
M adduct (95, 145). In stopped-flow experiments carried out on LL1 variants, we observed the 
spectral feature consistent with the presence of both compound I-like and compound-II like 
species. Thus, RFQ-EPR experiments were carried out on LL1 variants to identify and confirm 
the electronic nature of these two species and to observe the presence or absence of protein based 
radical in these variants. The data obtained from these experiments are summarized in chapter 3. 
2.7. Conclusions 
From this section, it is clear that wide array of instrumentations and techniques were 
required to study KatGs. These techniques ranged from molecular biology to various 
spectroscopic instrumentations. Molecular biology provided an opportunity to create new 
variants by genetic manipulation. In order to study those variants, specific expression and 
purification methods were employed to obtain the proteins in their native states. The structural 
impacts of manipulations were studied by spectroscopic techniques like UV-Vis, EPR etc., 
whereas appropriate kinetic characterizations were performed to observe the changes in proteins? 
activity and also to identify the intermediates formed in the catalytic mechanisms.  
 
88 
  
Chapter Three: Tracing the Impact of a Unique Loop in Catalase-peroxidase 
Catalysis 
3.1. Introduction 
 Catalase-peroxidases (KatGs) are heme-containing enzymes with a multifunctional active 
site. The enzymes are named after their catalase and peroxidase activities, but they also posses 
peroxynitritase (146), NADH oxidase, isonicotinoyl hydrazide INH lyase, and isonicotinoyl-
NAD synthase activities (89). Due to the latter two activities, KatG has been implicated as the 
enzyme responsible for the activation of a frontline antitubercular drug (isoniazid) and is, 
therefore, central to the treatment of  Mycobacterium tuberculosis infections (147, 148). 
Depending upon the geographical distribution, in 50-70% of isoniazid-resistance M. tuberculosis 
clinical isolates, katG gene is the most commonly targeted gene with 30-90% of the mutation 
occurring in codon 315 (149-152). However, neither the mechanism of isoniazid activation by M. 
tuberculosis KatG nor the mechanism by which mutations to the katG gene lead to resistance is 
fully understood. 
 The catalytic mechanism of KatG as well as the nature of the interplay between the two 
dominant activities, catalase and peroxidase, remain a matter of some discussion. It is generally 
agreed that catalase and peroxidase reactions involve a common path for the formation of a ferryl 
protoporphyrin IX (PPIX) ?-cation radical intermediate (i.e., compound I) but differ in the 
 
89 
  
mechanism for compound I reduction back to the ferric state. Many have argued that the catalase 
activity of KatG follows a mechanism similar to that of typical or monofunctional catalases. As 
such, there are two major steps. In the first step, the ferric enzyme undergoes a two-electron 
oxidation by one equivalent of H2O2, yielding compound I and H2O (reaction 3.1). In the second 
step, the enzyme returns to its resting state by reacting with a second equivalent of H2O2, this 
time oxidizing it to O2 (reaction 3.2). With typical enzymes from the non-animal peroxidase 
superfamily, compound I is returned to the ferric state by two single-electron reduction steps at 
the expense of an exogenous electron donor. In the first transfer, compound I is reduced, 
generating a ferryl intermediate (i.e., compound II) and the first equivalent of the corresponding 
substrate radical (reaction 3.3). Compound II then oxidizes the second equivalent of the 
substrate, generating a second radical and returning the enzyme to its ferric starting state 
(reaction 3.4).  
 
H 2 O 2 + F e
I I I
( P P I X ) H
2 O
+ F e
I V
= O ( P P I X    ) ( 3 . 1 )
H 2 O 2 + H 2 O   +   O 2 + ( 3 . 2 )
R H + R + F e
I V
= O ( P P I X ) ( H   ) ( 3 . 3 )
R H + H 2 O + ( 3 . 4 )
H 2 O 2 + H 2 O + F e
I I I
- O 2  ( P P I X ) ( 3 . 5 )
F e
I V
= O ( P P I X    ) F e
I I I
( P P I X )
F e
I V
= O ( P P I X    )
F e
I I I
( P P I X )R +
F e
I V
= O ( P P I X )
F e
I V
= O ( P P I X ) ( H   )
 
90 
  
Though the mechanisms of typical catalases and peroxidases have many similarities, they 
are distinct from one another in important ways. In peroxidases, compound II is an essential 
component of the catalytic cycle. With the typical monofunctional catalases, compound II is 
regarded as an inactive species with respect to catalytic O2 production. When H2O2 
concentrations are high and/or exogenous electron donor concentrations are low, peroxidase 
compound II typically reacts quite easily with H2O2 to form a FeIII-O2.- intermediate known as 
oxyperoxidase or compound III (reaction 3.5). Compound III is inactive with respect to the 
turnover of typical peroxidases and is commonly observed in these enzymes. It is also an inactive 
intermediate for catalases, but compound III is almost never observed with these enzymes (71, 
153-156).  
By any definition, KatG is a robust catalase. Their active sites produce turnover numbers 
(i.e., apparent kcat values) of around 5000 s-1. Apparent second-order rate constants of 106 M-1s-1 
are typical (157). What makes this kinetic performance striking is that the KatG active site bears 
no resemblance to the typical monofunctional catalase enzymes (e.g., E.coli catalase [HPII]). 
Consistent with its membership in the non-animal peroxidase superfamily, the KatG active site is 
superimposable on the other Class I members of the superfamily (56, 62, 91-94). These enzymes 
(and all Class II and Class III peroxidases as well) have little if any catalase activity. Indeed, 
typical kcat values for catalase turnover (which is referred to as pseudocatalase activity) are near 
10 s-1 (71, 154). Due to this strange structure/function relationship the very existence of the 
catalase-peroxidase raises fundamental questions about heme enzyme structure, catalysis, and 
evolution. How is an existing enzyme active site with an obvious catalytic function co-opted to 
accommodate an entirely new function? What does this tell us about how nature has approached 
 
91 
  
this shift of functional capability? The active sites have changed little between catalase-negative 
peroxidases and highly catalase active KatG. This would suggest that the structures external to 
the active site must have some obvious differences.  
Indeed, catalase-peroxidases do contain a group of structural components which are 
absent from their monofunctional counterparts. First, each catalase-peroxidase subunit contains a 
second domain; a gene-duplicated C-terminal domain which has lost its ability to bind heme or 
to catalyze any reaction of its own (71, 91-94, 99, 158). However, the Goodwin laboratory has 
demonstrated that the presence of this domain is necessary to maintain catalase-peroxidase 
structure and function (86, 100). In particular, the C-terminal domain supports the architecture of 
the active site by preventing coordination of the heme iron by so called the distal histidine. Also, 
the C-terminal domain, when it is expressed and isolated as a separate protein, can restore the 
function of the N-terminal domain (100). 
In addition, the active site-bearing N-terminal domain contains three insertions which are 
absent from typical peroxidases: Large Loop 1 (LL1), Large Loop 2 (LL2), and Large Loop 3 
(LL3) (94). LL3 is the smallest of the three and shows a great deal of sequence variability. As 
yet, no specific function has been proposed or assigned for this structure. LL1 and LL2 are 
highly conserved across all catalase-peroxidases, but in fundamentally different ways. LL2 
shows considerably more sequence variability than LL1, but its length is essentially invariant. 
Conversely, LL1 shows high degree of sequence conservation but the length of the structure is 
highly variable from one species to another. In spite of these differences, both structures have 
been shown to be essential for catalase but not peroxidase activity (87, 90, 108, 113). 
 
92 
  
The LL1 contains a strictly conserved sequence of ten amino acids (MGLIYVNPEG) at 
its C-terminal end. The tyrosine (Y226 by E. coli numbering) participates in a unique Trp-Tyr-
Met (W-Y-M) covalent adduct as observed in the crystal structures of all catalase-peroxidases 
structures solved to date (91-94). Formation of this covalent adduct is heme and peroxide 
dependent (108), and substitution of any of these three amino-acids results in the loss of catalase 
activity with either little effect on or enhancement of peroxidase activity (90, 95-97, 108-110, 
113). The redox cycling of the W-Y-M adduct between its fully covalent and one-electron 
oxidized states has been suggested to as one mechanism to facilitate the enzyme?s catalatic 
activity (144). According to this mechanism the free radical oxidized state of the W-Y-M adduct 
can serve as an electron acceptor what is typically regarded as the peroxidatic dead end 
intermediate (FeIII-O2?  ). The electron transfer from FeIII-O2?   to the  -Y-M radical would 
produce the FeIII-O2 complex which would immediately dissociates, yielding FeIII enzyme and 
O2. At the very least this could serve as a protective mechanism by preventing FeIII-O2?   
accumulation, and may account for the bulk of catalase activity observed in KatG (reactions 3.6-
3.9) (75, 117, 159). 
H 2 O 2 + F e I I I ( P P I X ) ( W Y M ) H 2 O + F e
I V = O ( P P I X    ) ( W Y M ) ( 3 . 6 )
( 3 . 7 )F e
I V = O ( P P I X    ) ( W Y M ) F e I V = O ( P P I X ) ( W Y M    )
H 2 O 2 + F e I V = O ( P P I X ) ( W Y M    ) H 2 O + F e
I I I - O
2  ( P P I X ) ( W Y M    ) ( 3 . 8 )
F e I I I - O 2  ( P P I X ) ( W Y M    ) F e I I I ( P P I X ) ( W Y M ) +   O
2
( 3 . 9 )
 
 
93 
  
Both LL1 and LL2 contribute to a narrow access channel to the deeply buried heme in the 
active site. The strictly conserved residues, Ile248, Asn251, and Glu253 (Synechocystis 
numbering), have been identified as essential for the H2O2 oxidation required for catalase 
turnover (91-94, 102). Substitution of any of these residues resulted in 50-70 % loss of catalase 
activity but retention of peroxidase activity. Ile248 maintains the heme architecture by forming a 
hydrogen bond with Asp152 (105), and Glu253 creates an acidic entrance to the access channel 
which stabilizes and orients the solute matrix essential for H2O2 oxidation (102). Even though 
the LL1 structure is 36-76 amino-acids long and it connects distal and proximal pocket of the 
active site (Figure 3.1), the dominant role which has been articulated for LL1 to this point is to 
position tyrosine correctly for the formation of W-Y-M adduct. However, this could be 
accomplished with a much less extensive structure and as such, it raises a question: Does this 
structure play other roles in regulating catalase and peroxidase activities?  In order to investigate 
this question, three deletion variants were created in E.coli KatG; KatG?LL1 (Leu193-Asn228), 
KatG?209-228 (Ser209-Asn228), and KatG?200-214 (Glu200-Lys214) and compared to wt KatG and 
the substitution variant Y226F KatG. These variants were created by keeping in mind that each 
deleted different portions of LL1 and only two of them contained the essential tyrosine (Y226). 
The effects of each deletion on the overall stability and catalytic activity of KatG were monitored 
by various biophysical techniques.  
 
94 
  
 
 
Figure 3.1: LL1 Connecting Distal and Proximal Part of the Active Site in KatG. Structure 
was taken from PDB accession domain 1SJ2. Different structures are color coded as follows 
Heme-red, Large loop 1 (LL1) ? Green and Blue. 
  
 
S 209
M 252
I 248
W 105
Y 226
D 377
N 228
H 267
W 318
 
95 
  
3.2 Materials and Methods 
3.2.1. Materials 
All oligonucelotide primers were obtained from Invitrogen (Carlsbad, CA). E. coli stains 
(XL-1 Blue and BL-21 [DE3]) and Pfu polymerase were purchased from Agilent Technologies 
(La Jolla, CA). The restriction enzymes were obtained from New England Biolabs (Beverly, 
MA). QIAprep Spin Miniprep kit and nickel-nitrilotriacetic acid (Ni-NTA) resin were obtained 
from Qiagen (Valencia,CA). Isopropyl-B-D-thioglactopyranoside (IPTG) was obtained from 
Gold Biotechnology Inc. (St. Louis, MO). Mono and di-basic sodium phosphate, acetic acid, and 
sodium acetate were purchased from Fisher (Pittsburgh,PA). Bugbuster and benzonase were 
purchased from EMD (Madison, WI). Ampicillin, chloramphenicol, imidazole, hemin, sodium 
dithionite, 2,2?-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) (ABTS), and hydrogen 
peroxide (30%), were obtained from Sigma (St. Louis, MO). Water purified through Barnstead 
Easy Pure II UV ultrapure water system (18.2 M?/cm resistivity) was used to make all buffers 
and media. 
3.2.2 Site-directed and deletion mutagenesis 
Y226F KatG variant was developed by site-directed mutagenesis using QuikChangeTM  
protocol as described in chapter 2. The primers used were ECCP m23 (+) (5?-GAG ATG GGT 
CTG ATC TTC GTT AAC CC-3?) and ECCP m23 (-) (5?-GGG TTA ACG AAG ATC AG CCC 
ATC TC-3?).  The nucleotide substitutions corresponding to the mutations are in bold italics. 
 
96 
  
PCR product was treated with Dpn I to remove starting template and ligated using T4 DNA 
liagase. The desired plasmids were used to transform E. coli XL-1 blue cells by electroporation. 
The amplicillin resistant colonies were screened by Hpa I digest as the successful mutants 
contain a unique Hpa I site. Positive colonies were submitted for full DNA sequence analysis to 
Davis Sequencing. Corrected mutated plasmids were used to transform E. coli BL21-
Gold[DE3]plysS. 
To generate constructs for the expression of KatG?LL1 (wtKatG lacking LL1 insertion), a 
deletion protocol based on Seamless?cloning was employed. The primers used were ECCPD101 
(CC AGC TCT TCC CGG ATC CGG TTC CCA GAC GTC TTC) and ECCPD102 (ATT TAC 
TCT TCC CCG GAA GC CCG GAT CAC AGC). The underlined sequence is the Eam 1104I 
recognition sequence and the ones in bold are the overhanging sequence remained after the 
hydrolysis with Eam 1104I. The desired plasmids were used to transform E. coli XL-1 blue cells 
by electroporation. Positive colonies were submitted for full DNA sequence analysis to Davis 
Sequencing. Corrected mutated plasmids were used to transform E. coli BL21-Gold[DE3]plysS. 
The variants KatG?200-214 and KatG?209-228 were generated by an alternate mutagenesis 
method due to the difficulty in obtaining methylated dCTP required for the Eam1104I-dependent 
protocol. The primers were designed by excluding corresponding codons for the deletion and 
they produced linear PCR product. This alternate procedures required modification of primers to 
include 5?- phosphoryl group for the blunt ligation of the PCR products. The primer pair for 
KatG?200-214 was: 5?-phosphate-GCA CCG CTG GGT GCA ACC GAG ATG GGT CTG-3? and 
5?-phospahte-ATC ACC CCA GTT AAC ATC CAG ATC CGG TTC-3?. For KatG?209-228 
 
97 
  
following primer pair was used: 5?-phosphate-CCG GAA GGC CCG GAT CAC AGC GGC 
GAA CCG-3? and 5?-phosphate-ATG ACG GTG AGT CAG CCA GGC TTT TTC ATC-3?. The 
desired plasmids were used to transform E. coli XL-1 blue cells by electroporation. Plasmids 
from candidate colonies were evaluated by diagnostic restriction digest and DNA sequence 
analysis. Corrected mutated plasmids were used to transform E. coli BL21-Gold[DE3]plysS. 
3.2.3. Expression and purification 
Y226F KatG, KatG?209-228, and KatG?200-214 were expressed in the soluble form whereas 
KatG?LL1 was expressed in inclusion bodies. Expressions of Y226F KatG, KatG?LL1, KatG?209-
228, and KatG?200-214 were carried out as described in chapter 2. Soluble purification protocol was 
used for Y226F KatG, KatG?209-228, and KatG?200-214 whereas insoluble purification was 
employed for KatG?LL1. Concentrations of purified enzyme was estimated according to the 
method of Gill and von Hippel (126) [?280(Y226F KatG) = 1.42 x 105 M-1cm-1 and ?280 (KatG?209-
228)=1.36 x 105 M-1cm-1, ?280 (KatG?200-214)=1.37 x 105 M-1cm-1, and ?280 (KatG?LL1)=1.31 x 105 
M-1cm-1].  
3.2.4. Absorption spectra and activity assays 
WtKatG and its variants were reconstituted with 0.9 equivalents of hemin. Following 
reconstitution, each protein was incubated for 24 hours at 4 C. The solution was then centrifuged 
to remove free hemin and other insoluble materials. The concentration of reconstituted enzyme 
was determined using the pyridine hemochrome assay (124). All spectra were obtained at room 
 
98 
  
temperature in 200 mM phosphate buffer, pH 7.0 using a Shmiadzu UV-1601 spectrophotometer 
(Columbia, MD) with a cell pathlength of 1.0 cm.  
Catalase activity was evaluated by monitoring the decrease in H2O2 concentration in the 
presence of 10 nm enzyme at 240 nm (?240= 39.4 M-1 cm-1) (135).  All assays were carried out at 
room temperature using Shimadzu UV-1601 spectrophotometer using 200 mM phosphate buffer, 
pH 7.0. Peroxidase activity was measured by monitoring the production of ABTS radical at 417 
nm (?417 = 34.7 mM-1 cm-1), initial concentration of ABTS was measured spectrophotometrically 
at 340 nm (?340 = 3.66 x 104 M-1 cm-1) (136). 
3.2.5. Rapid Freeze Quench (RFQ) EPR  
Rapid Freeze Quench (RFQ)-EPR experiments were performed in collaboration with Dr. 
Aimin Liu, Georgia State University. Solutions of wtKatG or Y226F KatG or KatG?209-228 (~260 
?M ) and varying equivalents (1, 5, 10, and 50) of H2O2 in 50 mM phosphate buffer, pH 7.0 were 
mixed in 1:1 ratio. The mixture was then incubated for the indicated time periods followed by 
freeze quench in isopentane at -130 C. The frozen samples were then packed in precision bore 
EPR tubes immersed in an isopentane bath. EPR tubes were then transferred to liquid nitrogen 
and finally examined using Bruker ER200D spectrometer with 9.6 GHz modulation frequency 
with a 4116DM resonator. For temperature control an oxford ITC 503S temperature controller, 
an ESR910 helium cryostat and LLT650/13 liquid helium transfer tube (Oxford Instrument, 
Concord, MA). Data acquisition software was developed in house and Bruker WINEPR V 2.11 
was used for data manipulation. 
 
99 
  
3.2.6. Stopped-flow analysis 
3.2.6.1 in the presence of H2O2 
In Single mixing experiments, Y226F KatG and KatG?209-228 was allowed to react with 
excess of H2O2 to form higher oxidation states. In this mode, syringe ?A? contained 6 ?M 
enzyme in 50 mM phosphate buffer, pH 7.0 and syringe ?B? contained varying concentrations of 
H2O2. Full spectra recorded by diode array indicated transient H2O2 concentration-dependent 
spectral shifts consistent with compound I formation followed by an H2O2-independent shift to a 
compound II-like spectrum. The rates of these two events provided a narrow H2O2 concentration 
range to measure a rate constant for compound I formation (5 to 30 equivalents or 7.5 ?M to 45 
?M H2O2). This rate was recorded at 412 nm using single wavelength detection. This wavelength 
allows more facile distinction between the formation and decay of the compound I intermediates. 
Similar results were obtained when absorption at 400 nm was monitored. 
3.6.2.2 in the presence of peroxidatic electron donors 
The return of ferryl heme intermediates to the ferric state in the presence of electron 
donors; ascorbate and ABTS, was evaluated using sequential mixing stopped-flow experiments.   
6 ?M of enzyme (Y226F KatG or KatG?209-228) was mixed against 0.95 equivalent of H2O2 an 
allowed to react in the delay line for 10 s or 0.7 s, respectively. These delay times allowed 
accumulation of compound II like intermediates because accumulation of a large fraction of 
compound I intermediate under single turnover conditions is near impossible. The contents of the 
delay line were mixed with 50 mM phosphate buffer, pH 7.0, as well as 30 ?M of ascorbate and 
 
100 
  
varying concentrations of ABTS. Thus, the final reaction mixture contained 1.5 ?M enzyme, 15 
?M ascorbate, 25 mM buffer, and 0-120 ?M ABTS. The full spectra were recorded by diode 
array, and the rate constants for ABTS-dependent reactions were measured at 400 nm by single 
wavelength measurements. Ascorbate reduces ABTS radicals as rapidly as they are generated to 
its parent compound ensuring a constant substrate concentration during the reaction (114, 139). 
Ascorbate is often a poor substrate for peroxidase enzymes and has been a very effective tool for 
measuring rapid peroxidase reactions with electron donors and minimizes the interference from 
the oxidized products. Moreover, the concentration of ascorbate was a constant in our ABTS 
stopped-flow studies making its contribution easy to account for. 
3.3 Results 
The importance of LL1 in positioning tyrosine 226 to participate in the W-Y-M covalent 
adduct (an essential feature for the catalase activity of KatG) has been extensively evaluated. 
Considerably less attention has been focused on other important roles which LL1 might be 
contributing in order to maintain the stability and functionality of KatGs.  
In an attempt to decipher what additional contributions LL1 may make to KatG function, 
we targeted LL1 by deleting portions of the structure including or excluding the essential 
tyrosine. In case of Y226F variant, substitution with phenylalanine eliminated the possibility of 
W-Y-M formation. KatG?LL1 variant was deleted by removing amino acids from Leu193 to 
Asn228, and also lacked the ability to form the covalent adduct. KatG?200-214 variant was created 
to leave out the Y226 by deleting Glu200-Lys214 at the N-terminal portion of LL1. The 
 
101 
  
KatG?209-228 variant removed Ser209-Asn228 at C-terminal part of LL1 and it did include the 
essential tyrosine. These variants were investigated by steady-state and transient-state kinetics.  
3.3.1. Steady-state kinetic analysis 
The variants KatG?LL1, KatG?209-228 and Y226F KatG exhibited no catalase activity 
whereas KatG?200-214 retained catalase activity comparable to the wtKatG (Table 3.1). The 
substitution of tyrosine or its absence in KatG?LL1, and KatG?209-228 is sufficient to explain the 
loss of catalase activity. As previously mentioned, this tyrosine is involved in covalent adduct 
required to maintain the catalase activity in wtKatG. It has been shown that the analogous 
replacement of tyrosine 226 with phenylalanine in KatG from several other organisms like M. 
tuberculosis, B. pseudomallei and Synechocystis eliminates the catalase activity (108, 113). 
Interestingly, the effect on peroxidase activity of tyrosine substitution of KatG from  these 
organisms varied considerably ranging from little effect on peroxidase activity (90) to substantial 
increase (108, 113) or decrease in peroxidase activity (89). Our evaluation of Y226F from E.coli 
KatG indicated that any of these three outcomes may be observed depending upon the conditions 
used to measure peroxidase activity. 
The dependence of the peroxidase activity of Y226F KatG, KatG?200-214 and wtKatG on 
H2O2 is shown in Figure 3.2 (Panel A). Y226F KatG showed substantially greater peroxidase 
activity than either wtKatG or KatG?200-214 provided that H2O2 concentrations were relatively 
low. The apparent second order rate constant (kcat/KM) for Y226F KatG (2.3 ? 106 M-1s-1) with 
respect to H2O2 was nearly one order of magnitude greater than that determined for wtKatG  
 
102 
  
 
 
 
 
 
Table 3.1: Catalase kinetic parameters for wild-type KatG, Y226F KatG, and deletion 
variants 
 
Protein   Kinetic Parameters 
  kcat (s-1) KM (mM) kcat/KM (M-1s-1) 
wtKatG 11000 ? 200 3.4 ? 0.1 3.2 ? 106 
Y226FKatG ND ND ND 
KatG?200-214 9600 ? 200 7.2 ? 0.2 1.3 ? 106 
KatG?209-228 ND ND ND 
KatG?LL1 ND ND ND 
N/D: Not detected  
 
 
 
 
 
 
103 
  
(3.6? 105 M-1s-1) (Table 3.2). This was due largely to a much lower apparent KM of Y226F KatG 
for H2O2. Indeed, the maximum observed peroxidase activity for Y226F KatG was only 1.4 fold 
higher than that recorded for wtKatG. Along with this, Y226F KatG and wtKatG showed starkly 
different sensitivities toward H2O2-dependent inhibition. At higher concentrations of H2O2 (i.e., 
those exceeding 0.8 mM) Y226F KatG showed a progressive and substantial loss of peroxidatic 
activity, but H2O2 produced a much more modest decrease in the peroxidase activity of wtKatG. 
The H2O2-depedent peroxidase activity of KatG?200-214 showed similar response to varying 
concentration H2O2 compared to wtKatG as can be observed from comparable apparent second 
order rate constants.  
The effects of H2O2 on the peroxidase activity of the two deletion variants KatG?LL1 and 
KatG?209-228 were compared to Y226F KatG (Figure 3.2, panel B). The apparent kcat values for 
KatG?209-228 and KatG?LL1 were 8- and 4-fold higher, respectively, when compared to that of 
Y226F KatG. Closer examination of kinetic data in Figure 3.2, panel B showed that these three 
variants depicted a very different response at low and high concentrations of H2O2. Even though, 
all the three variants showed similar response to lower concentrations of H2O2 (inset), they 
diverged dramatically at higher concentrations of H2O2 specifically in terms of their 
susceptibility to H2O2-dependent inhibition. Out of the three variants, KatG?209-228 showed least 
susceptibility to inhibition as indicated by the largest KI value (~10 mM) while Y226F KatG was 
the most susceptible variant with the lowest KI value (~0.7 mM). KatG?LL1 showed intermediate 
 
104 
  
 
. 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
Figure 3.2: Effect of H2O2 concentration on the peroxidase activity of wild-type KatG, 
Y226F KatG, and deletion variants (KatG?LL1, KatG?200-214, and KatG?209-228). Panel A 
compares the activities of wild-type KatG (?), KatG?200-214 ( ? ) and Y226FKatG (?). Panel B 
compares the activities of variants lacking observable catalase activities, these are KatG?209-228 
(?), KatG?LL1 (?), and Y226F KatG (?). Activities of catalase negative variants at low H2O2 
concentrations are shown in the inset. All reaction contained 0.5 mM ABTS, 50 mM acetate 
buffer, pH 5.0, and were carried out at 23?C. 
0 . 0 0 . 4 0 . 8 1 . 2 1 . 6 2 . 0
0
20
40
60
80
A
[H
2
O
2
]  m M
V
/
[
E
]
T
 (
s
-1
)
0 2 4 6 8 10
0
100
200
300
400
500
600
700
800
900
B
[H
2
O
2
] m M
V
/[E
]
T
 (
s
-1
)
0 .0 0 0 .0 5 0 .1 0 0 .1 5 0 .2 0
0
50
100
150
200
[H
2
O
2
]  m M
V
/
[E
]
T
 
(
s
-1
)
 
105 
  
 
 
 
 
 
 
Table 3.2: Peroxidase kinetic parameters with respect to H2O2 for wild-type KatG, Y226F 
KatG, and deletion variants 
 
 
Protein   Kinetic Parameters   
  kcat (s-1) KM (mM) kcat/KM (M-1s-1) KI (mM) 
wtKatG 76 ? 9 0.22 ? 0.05 3.5 ? 105 3 ? 1 
Y226F KatG 97 ? 5 0.043 ? 0.005 2.3 ? 106 0.73 ? 0.08 
KatG?200-214 50 ? 2 0.09 ? 0.01 5.6 ? 105 5 ? 1 
KatG?209-228 904 ? 34 0.83 ? 0.06 9.6 ? 105 10.0 ? 0.8 
KatG?LL1 713 ? 86 0.97 ? 0.17 7.4 ? 105 1.7 ? 0.3 
 
 
 
 
 
 
106 
  
sensitivity to inhibition with a KI value of around 1.7 mM. It is likely that at least one contributor 
to the disparity between maximum observed activities and apparent kcat values was the 
magnitude of the KI with respect to H2O2. 
The effect of reducing substrate (ABTS) on the peroxidase activity of KatG?209-228, 
KatG?LL1, and Y226F KatG was also evaluated (Figure.3.3). When compared to H2O2-dependent 
activities, even though the data obtained for ABTS did not fit particularly well to a hyperbolic 
equation, a similar trend in the overall peroxidase activities was observed with the greatest 
activity displaced by KatG?209-228 followed by KatG?LL1 and then Y226F KatG. However, in 
contrast to the H2O2?dependence where there was a different response to low and high 
concentrations of H2O2, ABTS-dependent activities of the variants maintained the same trend at 
all concentrations of ABTS tested (i.e., above and below the apparent KM for ABTS). 
3.3.2. UV-visible characterization 
 The UV-visible absorption spectra were recorded for the ferric form of Y226F KatG, and 
KatG?209-228 following reconstitution with hemin and compared with those recorded previously 
for wtKatG and KatG?LL1 (103). The ferric state of both the variants showed a Soret maximum at 
~407 nm and charge transfer maxima at 508 and 642 nm indicating a mixture of states dominated 
by pentacoordinate and hexacoordinate high-spin heme. All UV-visible features were 
comparable to wtKatG and KatG?LL1; except that KatG?LL1 showed a substantial contribution 
from hexacoordinate low-spin heme species (103). 
Upon addition of 100 ?M H2O2, the UV-Vis spectra of Y226F KatG and KatG?209-228 
 
107 
  
 
 
 
Figure 3.3: Effect of ABTS concentration on the peroxidase activities of the catalase 
negative variants KatG?209-228 (?), KatG?LL1 (?), and Y226F KatG (?). All reactions for 
Y226FKatG, and KatG?LL1 contained1 mM H2O2. All reactions for KatG?209-228 contained 2 mM 
H2O2. Regardless of variant all assays were performed using 50 mM acetate buffer, pH 5.0, and 
carried out at 23?C. 
 
0 . 0 0 . 2 0 . 4 0 . 6 0 . 8 1 . 0
0
100
200
300
400
500
600
700
[ A B T S]  m M
V
/[
E
]
T
(s
-1
)
 
108 
  
showed a red shift of the Soret band to 418 nm and emergence of ? and ? bands at 542 nm and 
578 nm, respectively. This suggested that variants lacking tyrosine 226 either by substitution or 
deletion both easily formed the ?inactive? compound III intermediate in the presence of H2O2 
(Figure.3.4 A and B). As described earlier, formation of inactive species (FeIII-O2?   ) or 
oxyperoxidase is a common mechanism by which H2O2 inhibits peroxidases. This species is 
formed when compound II (FeIV=O) reacts with H2O2 rather than the peroxidase reducing 
substrate. This formation of compound III with H2O2 in the absence of a reducing substrate has 
been reported in KatG enzymes from other organisms where the tyrosine analogous to tyrosine 
226 was substituted with phenylalanine (75, 95, 113, 142, 160).  
3.3.3. Transient-state kinetic analysis: 
3.3.3.1. Single mixing reaction with H2O2 
Though each of the variants formed compound III quite easily in the presence of 
relatively low concentration of H2O2 (100 ?M), the steady-state kinetic studies clearly showed a 
different susceptibility to inactivation for each variant. In order to investigate this further, 
stopped-flow kinetic measurements were carried out to monitor the effect of H2O2 on the rates of 
interconversion between heme intermediates.  
In general, KatG compound I is a difficult species to observe in the presence of H2O2 
because of its rapid reduction by H2O2 in the catalase cycle. However, in catalase-negative 
variants it is conceivable that the reaction may be observed. Expected spectral changes would 
include 40-50% hypochromicity of Soret band and exponential decrease in the absorbance over  
 
109 
  
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
Figure 3.4: Effect of H2O2 on the absorbance spectra of Y226F KatG (A), and KatG?209-228 
(B). Spectral changes were recorded before (dashed line) and after (sold line) addition of 100 ?M 
H2O2. All spectra were recorded using 50 mM phosphate buffer, pH 7.0, and at 25 ?C 
300 400 500 600 700
0 . 0
0 . 1
0 . 2
0 . 3
0 . 0
0 . 1
0 . 2
0 . 3
407 418
542
578
642
B
W a v e le n g t h  ( n m )
Abs
o
r
b
a
n
c
e
Abs
o
r
b
a
n
c
e300 400 500 600 700
0 . 0
0 . 1
0 . 2
0 . 3
0 . 0
0 . 1
0 . 2
0 . 3
407
418
542
578
642
A
W a v e le ng t h ( nm )
A
bs
o
r
ba
nce
A
bs
o
r
ba
nce
 
110 
  
time at a wavelength closer to Soret maximum.  hen 3 ?M of ferric Y226F KatG was reacted 
with 5 equivalents of H2O2, 20-30 % hypochromicity of Soret band was observed with two 
isosbestic points (341 and 420  nm) suggesting formation of a compound I-like species (Figure 
3.5).  Similar spectral changes were observed upon reaction of 3 ?M of ferric KatG?209-228 with 
10 equivalents of H2O2 (Figure 3.6). In each case, the transition was monophasic as observed at 
400 nm (see Figure 3.5 and 3.6 insets) indicating pseudo-first order conditions had been 
achieved. A plot of kobs versus H2O2 concentration was linear for both the variants. However, the 
second order rate constant for the formation of the compound I-like species was two-fold higher 
for Y226F KatG  (7.5 ? 106 M-1s-1 ) compared to KatG?209-228 (3.9 ? 106 M-1s-1) (Figure.3.7).  
The compound I-like species of both the variants was only transiently observed, and in 
the absence of an exogenous electron donor an intermediate with spectral features suggestive of 
compound II-like species appeared. This intermediate was characterized by Soret band at 418 nm 
and ? and ? bands at 530 and 560 nm, respectively. It was formed within ~0.045 s for Y226F 
KatG (Figure 3.8) and ~0.05 s for KatG?209-228 (Figure 3.9).  In reaction traces recorded at 412 
nm, the formation of compound II-like intermediate comprised the second phase of a biphasic 
transition. The first phase corresponded to formation of compound I-like species followed by its 
decay and the formation of a compound-II like species in the second phase (insets, Figures 3.8 
and 3.9).  
The dependence of kobs for the second phase on H2O2 concentration was evaluated. There 
was a hyperbolic response to H2O2 for Y226F KatG and and KatG?209-228. Overall, rates were 
more rapid for Y226F KatG than KatG?209-228 with the former approaching a maximum for a first  
 
111 
  
 
 
 
 
 
 
 
Figure 3.5: Compound I formation of Y226F KatG. Spectral changes observed upon addition 
of 15 ?M H2O2 to 3 ?M ferric Y226F. First spectrum is that of ferric enzyme. Following spectra 
were taken after 2.5, 3.8, 6.3 and 15 ms (bold), respectively. Reactions were carried out in 50 
mM phosphate buffer, pH 7.0 and 4 ?C. Typical time trace between 3 ?M Y226F and 15 ?M 
H2O2 followed at 400nm shown in inset. 
 
 
 
300 400 500 600 700
0 . 0 0
0 . 0 4
0 . 0 8
0 . 1 2
0 . 1 6
0 . 2 0
0 . 2 4
0 . 0 0
0 . 0 4
0 . 0 8
0 . 1 2
0 . 1 6
0 . 2 0
0 . 2 4
407
510
W a v e le n g t h  ( n m )
A
b
s
o
r
b
a
n
c
e
A
b
s
o
r
b
a
n
c
e
0 . 0 0 . 1 0 . 2
0 . 1 6
0 . 1 8
0 . 2 0
0 . 2 2
T i m e  ( s )
A
400
 
112 
  
 
 
 
Figure 3.6: Compound I formation of KatG?209-228. Spectral changes observed upon addition 
of 15 ?M H2O2 to 3 ?M ferric KatG?209-228. First spectrum is that of ferric enzyme. Following 
spectra were taken after 2.5, 3.8, 5 and 13 ms (bold), respectively. Typical time trace between 3 
?M KatG?209-228 and 15 ?M H2O2 followed at 400nm is shown in inset. Reactions were carried 
out in 50 mM phosphate buffer, pH 7.0 and 4 ?C. 
 
 
 
300 400 500 600 700
0 . 0 0
0 . 0 5
0 . 1 0
0 . 1 5
0 . 2 0
0 . 2 5
0 . 3 0
0 . 0 0
0 . 0 5
0 . 1 0
0 . 1 5
0 . 2 0
0 . 2 5
0 . 3 0
407
510
W a v e leng t h  ( n m )
Abs
o
r
b
a
n
c
e
Abs
o
r
b
a
n
c
e
0. 0 0. 1 0. 2
0. 16
0. 17
0. 18
0. 19
0. 20
T i m e  ( s )
A
400
 
113 
  
 
 
 
 
 
Figure 3.7: Rate constants for formation of compound I in Y226F KatG (?) and KatG?209-
228 (?) plotted against H2O2 concentration at 400 nm. Reactions were carried out using 3?M 
of enzymes in 50 mM phosphate buffer, pH 7.0 at 4 C 
 
0 5 10 15 20 25 30
0
50
100
150
200
250
[H
2
O
2
] ( ? M)
k
o
b
s
(s
-1
)
 
114 
  
 
 
 
 
 Figure 3.8: Compound I decay to compound II of Y226F KatG. Spectral changes observed 
upon addition of 15 ?M H2O2 to 3 ?M ferric Y226F. First spectrum is that of compound I taken 
at 15 ms. Following spectra were taken after 45, 78, 127, 500, and 620 ms (bold, peaks at 418, 
530, and 560 nm, respectively). Reactions were carried out in 50 mM phosphate buffer, pH 7.0 
and 4 ?C. Typical time trace between 3 ?M Y226F and 15 ?M H2O2 followed at 412 nm showing 
decay of compound I to compound II is shown in inset. 
 
300 400 500 600 700
0 . 0 0
0 . 0 6
0 . 1 2
0 . 1 8
0 . 2 4
0 . 3 0
0 . 0 0
0 . 0 6
0 . 1 2
0 . 1 8
0 . 2 4
0 . 3 0
418
530
560
W a v e le n g t h  ( n m )
Abs
o
r
b
a
n
c
e
Abs
o
r
b
a
n
c
e
0 . 0 0 . 2 0 . 4
0 . 1 8
0 . 1 9
0 . 2 0
0 . 2 1
T i m e  ( s )
A
412
 
115 
  
 
 
 
 
 
 
 
Figure 3.9: Compound I decay to compound II of KatG?209-228. Spectral changes observed 
upon addition of 15 ?M H2O2 to 3 ?M ferric KatG?209-228. First spectrum is that of compound I 
taken at 13 ms. Following spectra were taken after 50, 75, 126, and 500 ms (bold, peaks at 418, 
530, and 560 nm, respectively). Reactions were carried out in 50 mM phosphate buffer, pH 7.0 
and 4 ?C. Typical time trace between 3 ?M KatG?209-228 and 15 ?M H2O2 followed at 412 nm 
showing decay of compound I to compound II is shown in inset. 
330 430 530 630
0 . 0
0 . 1
0 . 2
0 . 3
0 . 4
0 . 0 0
0 . 0 5
0 . 1 0
0 . 1 5
0 . 2 0
0 . 2 5
418
530
560
0 . 0 1 2 6
642
W a v el en g t h  ( n m )
A
b
s
o
rb
a
n
ce
A
b
s
o
rb
a
n
ce
0 . 0 0 . 1 0 . 2 0 . 3 0 . 4 0 . 5
0 . 1 9
0 . 2 0
0 . 2 1
0 . 2 2
0 . 2 3
T i m e  ( s )
A
412
 
116 
  
order rate constant of 23 ? 2s-1 while the latter approached 15 ? 1 s-1 (Figure 3.10). 
One interpretation consistent with the data would be that of two sequential reactions. The 
first is a bimolecular reaction with H2O2 to form a compound I-like species. The rate constant 
corresponding to this step is 7.5 or 3.9 ? 106 M-1s-1 for Y226F KatG and KatG?209-228, 
respectively. The second is an intramolecular electron transfer resulting in radical transfer from 
the heme to the KatG protein. The first order rate constant corresponding to this step being 23 ? 
2 s-1 for Y226F KatG and 15 ? 1 s-1 for KatG?209-228 (reaction 3.10)  
F e I I I ( P P I X ) ( P ) H 2 O 2 F e I V = O ( P P I X    ) ( P ) F e I V = O ( P P I X ) ( P  )k 1 k 27 . 5  ? 1 0 6  M - 1 s - 1
3 . 9  ? 1 0 6  M - 1 s - 1
2 3  ?  2  s - 1  
1 5  ?  1  s - 1
+ ( 3 . 1 0 )
 
Given the rate constants for each step, and the concentrations of H2O2 utilized, one would 
predict based on this model a change in rate determining step for the formation of the compound 
II-like species. At low H2O2 concentrations (less than 15 ?M) the first step would be rate 
determining, none of the compound I-like species would accumulate, and the reaction would 
appear as a direct conversion to the compound II-like species. Conversely, at concentrations of 
H2O2 15 ?M and above the rate of compound I formation would begin to exceed that of its 
conversion to the compound II-like species, and compound I would accommodate. Absorbance 
at 412 nm allows for monitoring of both steps. The effect of H2O2 on the character of the trace at 
412 nm is precisely consistent with the mechanism proposed and the rate constants determined. 
At the lowest H2O2 concentration tested for both variants a single phase conversion was 
observed. As H2O2 concentration increased, the biphasic character of the trace increased with the 
 
117 
  
 
 
 
Figure 3.10: Rate constants for formation of compound II in Y226F KatG (?) and 
KatG?209-228 (?) plotted against H2O2 concentration at 412 nm. Reactions were carried out 
using 3?M of enzymes, 50 mM phosphate buffer, pH 7.0 at 4 C. 
 
 
 
 
0 10 20 30 40 50 60
0
5
10
15
20
25
[H
2
O
2
] ( ? M)
k
o
b
s
(s
-1
)
 
118 
  
pronounced decrease in absorbance followed by a subsequent increase in absorbance became 
more pronounced. The first phase of the reaction was clearly H2O2 dependent, just as the second-
phase was H2O2 independent (Figure 3.11) 
As mentioned earlier, both KatG?209-228 and Y226F KatG reacted with 100 ?M H2O2 to 
rather easily form an intermediate with spectral characteristics consistent with FeIII-O2?   
intermediate (compound III), in order to determine the relative rates of the formation of this 
species, we monitored reactions with 100 ?M H2O2 by stopped-flow. Following accumulation of 
compound II-like intermediate a slow emergence of the FeIII-O2?   intermediate was observed 
indicated by a shift in the ? and ? bands from 530 and 560 nm, respectively to 542 and 578 nm, 
respectively (Figure 3.12 panels A and B). Interestingly, the rate of compound III formation was 
more rapid with KatG?209-228 thanY226F KatG. 
Both Y226F KatG and KatG?209-228 displayed similar mechanistic response to H2O2, and 
interestingly, the rate constants governing their reactions with H2O2 were very similar to one 
another. This would suggest that the profound differences between the two proteins in sensitivity 
to H2O2-dependent inactivation was not actually due to differences in the direct reaction with 
H2O2 or their relative rates. 
3.3.3.2. Rapid Freeze-quench EPR 
RFQ-EPR experiments were performed on Y226F KatG and KatG?209-228 to determine the 
presence of protein-based radicals during the reaction of these with H2O2. These studies were to  
 
119 
  
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
Figure 3.11: Typical time trace at 412 nm showing decay of compound I to compound II. 
Reaction of 3 ?M of either Y226F KatG (A) or KatG?209-228 (B) at different concentrations (?M) 
of H2O2 C, Overlay of the time traces of reaction of 3 ?M of Y226F KatG with15 ?M H2O2 
(solid line) and 3 ?M KatG?209-228 with 30 ?M H2O2 (dashed line). Reactions were carried out in 
50 mM phosphate buffer, pH 7.0 and 4 ?C. 
0 . 0 0 . 1 0 . 2 0 . 3 0 . 4 0 . 5
0 . 1 9
0 . 2 0
0 . 2 1
0 . 2 2
0 . 2 3
0 . 2 4
5
10
20
50
100
1
2
A
T i m e  ( s )
A
bs
 
(
4
1
2
 
nm
)
0 . 0 0 . 1 0 . 2 0 . 3 0 . 4 0 . 5
0 . 2 0
0 . 2 1
0 . 2 2
0 . 2 3
0 . 2 4
0 . 2 5
0 . 2 6
0 . 2 7
0 . 2 8
1
2
5
10
20
50
100
B
T im e ( s )
A
b
s
 
(
4
1
2
 
n
m
)
0 . 0 0 . 2 0 . 4 0 . 6 0 . 8 1 . 0
0 . 8 5
0 . 9 0
0 . 9 5
1 . 0 0
T im e  ( s )
A
bs
 
(
4
1
2
 
nm
)
C
 
120 
  
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
Figure 3.12: Heme intermediates observed during reactions with excess H2O2. Spectral 
changes upon addition of 100?M H2O2 to ferric Y226F (A) and KatG?209-228 (B) recorded at 0.09 
s (dashed line) and 7.5 s (solid line) for Y226F KatG and 0.2 s (dashed line) and 7 s (solid line) 
for KatG?209-228. C, Time course of reaction of Y226F KatG (dashed line) and KatG?209-228 (solid 
line) with H2O2 followed at 418 nm. All reactions were carried in 50 mM phosphate buffer, pH 
7.0 at 4 C  
300 400 500 600 700
0 . 0 0
0 . 0 4
0 . 0 8
0 . 1 2
0 . 1 6
0 . 2 0
0 . 2 4
0 . 0 0
0 . 0 4
0 . 0 8
0 . 1 2
0 . 1 6
0 . 2 0
0 . 2 4
418
542
578
530
560
A
W a v e len g t h  ( n m )
Ab
s
o
r
b
a
n
c
e
Ab
s
o
r
b
a
n
c
e
300 400 500 600 700
0 .0 0
0 .0 4
0 .0 8
0 .1 2
0 .1 6
0 .2 0
0 .2 4
0 .2 8
0 .0 0
0 .0 4
0 .0 8
0 .1 2
0 .1 6
0 .2 0
0 .2 4
0 .2 8
418
542
578
530
560
B
W a v e l e ng t h ( nm)
A
bs
o
r
ba
nc
e
A
bs
o
r
ba
nc
e
0 .0 0 .1 0 .2 0 .3 0 .4 0 .5
0 .6 5
0 .7 5
0 .8 5
0 .9 5
1 .0 5
2 4 6 8
T i m e  ( s )
R
e
l
a
t
i
v
e
 
A
bs
 
(
4
1
8
 
nm
) C
Fe
IV
=O
Fe
III
-O
2
.-
 
121 
  
complement the optical data for heme intermediates observed in our stopped-fow experiments. 
Both the variants were mixed with H2O2 and samples were frozen at an incubation times from 10 
ms to 400 ms. These conditions corresponded well  to optical stopped-flow measurements used 
to generate compound I and compound II (Figure 3.11).  
Figure 3.13 shows the results obtained when KatG?209-228 was mixed with 5 equivalents of 
H2O2 and frozen at the incubation times from 10 ms to 400 ms. At 5 equivalents of H2O2 an 
initial doublet with hyperfine splitting at ~21 gauss and line width of ~30 gauss was observed 
only after 10 ms (Trace A). On a longer time scale (300 ms) a singlet signal started to appear 
with much smaller hyperfine interaction and disappearance of doublet species (Trace B). At ten-
fold higher H2O2 concentration (50 equivalents) and a longer reaction time (400 ms) only a 
narrow singlet with no hyperfine splitting was observed (Trace C).  
The effect of reaction time and H2O2 concentration on radical species observed may 
reflect transition from the compound I-like to compound II-like spectra observed in our optical 
stopped-flow experiments. Moreover, the spectra clearly indicate the participation of protein 
components in oxidation even though the Tyrosine 226 side chain is absent and the W-Y-M 
covalent adduct cannot form. The Y226F variant presented substantially greater difficulty in the 
RFQ experiment.  Nevertheless, qualitatively similar results were obtained when the protein was 
mixed with 10 equivalents of H2O2 and allowed to react for 13 ms (Figure 3.14). 
3.3.3.3. Sequential mixing reaction with H2O2 and electron donors 
Single mixing kinetic experiments showed a slower rate of compound I formation and a 
faster rate of compound III formation for KatG?209-228 in the presence of H2O2 alone, indicating 
 
122 
  
 
 
 
 
Figure 3.13: EPR spectra of appearance of different species in KatG?209-228 during the 
reaction of resting enzyme with H2O2. KatG?209-228 (141 ?M) was reacted at 4 C with 5 
equivalents (A and B) and 50 equivalents of H2O2 (C). Reaction mixtures were freeze quenched 
after (A) 10 ms; (B) 300 ms; and (C) 400 ms. 
 
3300 3350 3400 3450 3500 3550
A
B
C
M a g net ic F ield ( G a us s )
 
123 
  
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
Figure 3.14: EPR spectra of appearance of doublet species in Y226F KatG during the 
reaction of resting enzyme with H2O2. Y226F KatG (131 ?M) was reacted at 4 C with 10 
equivalents of H2O2 and the reaction mixture was quenched after 13 ms. 
 
3350 3400 3450 3500
M a g n et i c F i eld  ( G a u s s )
 
124 
  
lower peroxidase activity and greater susceptibility to H2O2-dependent inactivation. However, 
substantially higher peroxidase activity and greater resistance to H2O2-dependent inactivation 
was observed for KatG?209-228. In order to account for these paradoxical findings, the reactions of  
KatG?209-228 and Y226F KatG with electron donors were evaluated by stopped-flow experiments. 
In order to carry out these experiments a sequential mixing approach was required. In this, both 
the enzymes were first reacted with single equivalent of H2O2, and then following mixing with 
solution containing electron donor, the return of ferryl heme intermediates to the ferric state was 
followed. In order to allow for formation of higher oxidation heme states in the initial reaction 
with H2O2, a delay time of 0.7 s and 10 s for Y226F KatG and KatG?209-228, respectively, was set 
to separate the initial mixing event and the final mixing event.  
For both the variants, the first spectrum observed under these conditions was the ferryl 
species resembling compound II. Reduction of compound II to the ferric state in the presence of 
buffer alone, ascorbate, or ascorbate and ABTS was followed by monitoring the absorbance 
increase at 400 nm (Figure 3.15, Panels A and B). Irrespective of the identity of electron donor, 
the rate of conversion of KatG?209-228   to the ferric state was substantially faster than Y226F 
KatG (Figure 3.15, Panel C). For the experiment including ascorbate and ABTS, we also 
evaluated the effect of ABTS concentration on the rate of enzyme reduction. The plot of pseudo 
first-order rate constants, kobs, against ABTS concentration was linear and from the slope second 
order rate constants were determined to be  5.2 ? 104 M-1 s-1 for KatG?209-228, which was over an 
order of magnitude higher than that for Y226F KatG (1.8 ? 103 M-1 s-1) (Figure3.16). 
 
125 
  
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
Figure 3.15: Reduction of compound II of Y226F KatG (A) and KatG?209-228 (B). Sequential 
mixing experiment were performed to monitor the return of ferryl intermediate to the ferric state 
in the presence of buffer (gray line), 15 ?M ascorbate (dotted line), and 15 ?M ascorbate and 15 
?M ABTS (solid line) under single turnover conditions. C, Effect of ABTS on the reduction of 
compound II of Y226F KatG (dotted line) and KatG?209-228 (solid line) to the ferric state. All 
reactions were carried out in 50 mM phosphate buffer, pH 7.0 at 4 C  
0 10 20 30 40
0 . 1 0
0 . 1 1
0 . 1 2
0 . 1 3
0 . 1 4
A
T i m e  ( s )
A
400
0 10 20 30 40
0 . 0 8
0 . 1 0
0 . 1 2
0 . 1 4
0 . 1 6
B
T i m e  ( s )
A
400
0 5 10 15 20 25 30 35 40 45
0 . 1 0
0 . 1 1
0 . 1 2
0 . 1 3
0 . 1 4
0 . 1 5
C
T im e  ( s )
A
400
 
126 
  
 
 
 
 
 
 
Figure 3.16: Rate constants for reaction of Y226F KatG (?) and KatG?209-228 (?) to the 
ferric state plotted against ABTS concentration. Effect of ABTS concentration, in the 
presence of 15?M ascorbate, on reduction of ferryl intermediates of each enzyme was monitored 
using sequential mixing stopped-flow experiment. All the reactions were carried out in 50 mM 
phosphate buffer, pH 7.0 at 4 C  
 
 
0 10 20 30 40 50 60
0 . 0
0 . 5
1 . 0
1 . 5
2 . 0
2 . 5
3 . 0
[ AB T S ]  ( ? M)
k
o
b
s
(s
-1
)
 
127 
  
3.4. Discussion 
Comparison of crystal structures and amino-acid sequences of catalase-peroxidases with 
their monofunctional peroxidase counterparts indicate the presence of a large loop 1 (i.e. LL1) 
between Asp192 and Pro229 (E.coli numbering). The LL1 structure forms part of substrate 
access channel and connects the distal and proximal halves of KatG active site. LL1 has been 
assigned an important role of supporting tyrosine (Y226) for the formation of Trp-Tyr-Met 
crosslink. This crosslink is an important feature that maintains the catalatic activity of KatG. 
However, the role of rest of the LL1 structure in KatG catalysis is not well defined. 
In this research, we have presented evidences in support of other important roles of LL1 
besides anchoring Y226 in its proper position. The variants involving Y226 have been created 
either by site-directed mutagenesis (Y226F KatG) or deletion mutagenesis (KatG?209-228 and 
KatG?LL1) and these were evaluated by steady state and transient-state kinetic methods. The 
properties of these variants were compared against the wtKatG and another LL1 deletion variants 
which retains tyrosine. Our data indicated that the LL1 structure restricts the access of 
peroxidatic electron donors to the active site and also that at least part of LL1 is important for 
maintaining the structural integrity of KatG. 
The importance of LL1 structure in stability of KatG protein was illustrated by 
comparison of KatG?LL1 to the variants where only part of the structure has been removed 
(KatG?209-228). KatG?LL1 was expressed in inclusion bodies and required purification under 
denaturing conditions followed by refolding, whereas KatG?209-228 was expressed in soluble 
form. Also, heme titration of apo KatG?LL1 produced a linear increase in activity only up to 0.25 
 
128 
  
equivalents, whereas wtKatG and KatG?209-228 showed no defects in heme uptake. As in all 
available crystal structures for KatG enzymes, the N-terminal and C-terminal ends of LL1 are 
separated by a considerable distance (~25 ?) (91-94). This instability in the KatG structure can 
arise due to the strain imposed on the enzyme structure by forcing D192 and P229 to be 
sequentially and therefore, spatially adjacent to one another. When a portion of the LL1 structure 
which does not appear to have a particular catalytic function (note the properties of KatG?200-214) 
is allowed to remain in the KatG structure as in KatG?209-228 the stability issues are largely 
resolved. 
Steady-state data showed that the LL1 variants involving Y226 either by deletion 
(KatG?LL1 and KatG?209-228) or substitution (Y226F) retained no catalase activity due to absence 
of the W-Y-M adduct, however, they all differed substantially in their peroxidase activity. The 
main effect on peroxidase activity was seen with respect to H2O2. These variants showed similar 
response to lower H2O2 concentrations, however, at higher H2O2 concentrations they diverged 
dramatically specifically in terms of their susceptibility to H2O2-dependent inhibition. The 
increase in peroxidase activity at lower H2O2 concentration can suggest that in the absence of 
catalatic activity, the catalytic machinery of KatG is completely dedicated towards 
accomplishing the peroxidatic activity of KatG. Furthermore, the inhibition at higher 
concentrations of H2O2 is suggestive of formation of an inactive intermediate FeIII-O2?   
(compound III).  
Apparent second-order rate constants for peroxidase turnover with respect to H2O2 were 
similar for each variant but with respect to the reducing substrate, ABTS, reactivity at low 
 
129 
  
concentrations varied considerably. The apparent second-order rate constant for Y226F KatG 
(1.7 ? 105 M-1s-1) was an order of magnitude lower than either KatG?209-228 (3.0 ? 106 M-1s-1) or 
KatG?LL1 (1.5 ? 106 M-1s-1).  
Single-mixing stopped-flow data showed that even though Y226F KatG showed poor 
peroxidase activity as compared to KatG?209-228, the rate of formation of compound II was twice 
as high for Y226F KatG than for KatG?209-228. Compound II is inactive in the catalase cycle but 
active in peroxidase cycle and its presence enhances the formation of compound III in the 
presence of H2O2. Indeed, both variants showed different susceptibility to peroxide-dependent 
inhibition via compound III formation. However, stopped-flow data indicate that the rate of 
accumulation of this intermediate is similar in both the variants.   
As the ferryl intermediate is able to react either with H2O2 forming compound III or 
ABTS leading to ferric state, a poor ability to accomplish latter would be expected to more facile 
accumulation of the FeIII-O2?   state. In some respect this is reminiscent of the structural features 
of typical monofunctional catalase enzyme which have a long and narrow channel leading to 
heme limiting peroxidase activity. Indeed, in S. cerevisiae catalase substitution intended to widen 
the access channel have been shown to increase the peroxidase activity (161). Sequential mixing 
stopped-flow studies confirmed that ferryl heme intermediates specifically compound II like 
species were rapidly converted to the resting ferric state in the presence of ABTS. Among the 
two variants, the rate to return to the ferric state was an order of magnitude faster for KatG?209-228 
than for Y226F KatG. 
 
130 
  
Figure 3.17 represents the proposed intermediates for the catalase (inner pathway) and 
peroxidase (outer pathway) activities of wtKatG. The catalase reaction in wtKatG incorporates a 
radical on W-Y-M adduct. The reaction of formation W-Y-M radical is proposed to proceed via 
formation of compound I (porphyrin ?-cation radical). This intermediate is rapidly converted to 
oxyferryl heme with simultaneous formation of W-Y-M radical (compound I*). The reaction of 
compound I* with second molecule of H2O2 produces an inactive intermediate (FeIII-O2?  ). 
Electron transfer from the superoxide radical of FeIII-O2?   to the  YM helps in decomposition of 
this intermediate to ferric enzyme and dioxygen (O2). In the peroxidase cycle, formation of an 
external protein radical (P?) is suggested from compound I?, keeping in mind the narrowness of 
the access channel in KatG compared to the size of many peroxidatic electron donors (AH). The 
intermediate carrying the P? was called compound I**. Compound I** was then reduced by two 
equivalents of AH generating ferric enzyme, two substrate radicals (A?) and a water molecule.  
Figure 3.18 proposes the modifications in the catalytic cycle of wtKatG in the absence of 
tyrosine, either by mutation (Y226F) or by deletion (KatG?LL1 and KatG?209-228). The absence of 
tyrosine prevents the formation of W-Y-M covalent adduct and this results in loss of catalase 
activity with variable effects on the peroxidase activity. In the absence of W-Y-M adduct, radical 
is proposed to transfer to external protein site generating P? (compound I**). Due to formation of 
P?, the second reaction with H2O2 would form FeIII-O2?   intermediate (inner dead end in Figure 
3.18). The easy formation of FeIII-O2?   in these variants, in the presence of low H2O2 
concentration, was supported by our UV-Vis and stopped-flow data (Figures 3.4 and 3.12). Also, 
the variable effects on peroxidase activity in Y226F KatG can be seen due to the barrier imposed 
by rest of the LL1 which limits the ability of exogenous electron donors to reach the active site. 
 
131 
  
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
Figure 3.17: Proposed intermediates for catalatic (inner pathway) and peroxidatic (outer 
pathway) activities of wild-type KatG. 
F e
I I I
P
W Y M
F e
I V
P
W Y M
F e
I I I
P
W Y M
O
F e
I V
P
W Y M
O
F e
I I I
P
W Y M
O
2
F e
I V
P
W Y M
O
F e
I V
P
W Y M
O
A H
A
H
2
O
H
2
OH
2
O
2
H
2
O H
2
O
2
A H
A
O
2
C o m p o u n d  I
C o m p o u n d  I *
C o m p o u n d  I * *
C o m p o u n d  I I
C o m p o u n d  I I I
N a t i v e
 
132 
  
 
 
 
 
 
 
 
 
 
 
 
 
.    
 
 
 
 
Figure 3.18: Proposed intermediates formed during the interrupted catalatic (inner dead 
end) and peroxidatic (outer pathway) activities of Y226F KatG and LL1 deletion variants. 
F e
I I I
P
W  M
F e
I V
P
W  M
F e
I I I
P
W  M
O
F e
I V
P
W  M
O
F e
I I I
P
W  M
O
2
F e
I V
P
W  M
O
F e
I V
P
W  M
O
A H
A
H
2
O
H
2
OH
2
O
2
H
2
O H
2
O
2
A H
A
2 A H
2 A  +  H
2
O
C o m p o u n d  I
C o m p o u n d  I * *
C o m p o u n d  I I I
C o m p o u n d  I * *
C o m p o u n d  I I
N a t i v e
 
133 
  
 This limited peroxidatic turnover can also favor the accumulation of FeIII-O2?   intermediate due 
to reaction with H2O2. However, the deletion variants in spite of being catalase negative due to 
the absence of W-Y-M adduct showed substantial greater peroxidase activity than Y226F KatG. 
An order of magnitude increase in the rate of return of ferryl species to ferric form in the deletion 
variants compared to Y226F KatG may suggest that deletion of entire LL1 or part of it relives 
the barrier and allows the access of larger exogenous electron donor leading to more direct 
electron transfer pathway (represented by bold dotted lines in Figure 3.18). This direct electron 
transfer increases the peroxidatic turnover in the deletion variants and ultimately prevents the 
accumulation of FeIII-O2?   intermediate. 
  In conclusion, we have shown that, LL1 act as a gate keeper to limit the peroxidatic 
activity in favor of catalase turnover. However, by obstructing the access of peroxidatic electron 
donor LL1 makes the enzyme prone to accumulation of inactive FeIII-O2?   intermediate. Under 
normal circumstances, this would be profoundly disadvantageous; however, presence of Y226 in 
W-Y-M in close proximity to heme takes advantage of FeIII-O2?   intermediate as a mechanism 
for release of oxygen and completion of novel catalatic mechanism. 
 
134 
  
Chapter Four: Dependence of Catalytic Ability of Catalase-peroxidases on 
Intersubunit Interactions 
4.1. Introduction 
 Catalase-peroxidases (KatGs) are heme-dependent, bifunctional enzymes found in 
bacteria, archebacteria and some fungi (21).  The catalase and peroxidase activities of these 
enzymes are similar to those of the robust monofunctional representatives found in nature.  The 
catalase activity of KatG rivals that of bovine liver catalase (32), and the peroxidase activity is 
well within the range of those observed within the non-animal peroxidase superfamily (105). 
This presents three options for the advent of this unique bifunctionality: 1) Enhanced peroxidase 
function has been built into the framework of a pre-existing catalase active site, 2) enhanced 
catalase activity has been obtained from a peroxidase active site , or 3) a completely new 
structural scaffold has been utilized which allows for both functions.  
The KatG active site is nearly identical to Class I of the non-animal peroxidases (e.g., 
cytochrome c peroxidase), but it has no resemblance to typical monofunctional catalases (98). 
Clearly, nature has taken the second option, modifying peroxidase scaffold to perform catalase 
activity orders of magnitude greater than any other members of its superfamily. The 
modifications in the peroxidase scaffold are immediately evident from the crystal structures and 
 
135 
  
amino acid sequence alignments of known KatGs in comparison to monofunctional peroxidases 
(91-94). On one hand KatGs are multimeric enzymes either homodimers or homotetramers. This 
is in contrast to most other non-animal peroxidases which are almost entirely monomeric. A 
direct monomer comparison reveals that the KatG subunit has three peripheral structures which 
are absent from the other monomeric peroxidases (91-94, 99) .  
 First, there is the gene duplicated C-terminal domain ? 30 ? away from the active site. 
Despite its substantial separation from the active site, it is essential for catalase-peroxidase 
function by preventing distal histidine coordination to the iron in the heme (86).  Second, two  
large loops, called large loop 1 (LL1) and large loop 2 (LL2), are observed in all the KatGs and 
have been shown to be essential for catalase but not peroxidase activity (43, 87, 98, 102, 162).  
Parts of LL1 and LL2 contribute to a narrow access channel to the deeply buried heme in the 
active site (103). In terms of length and sequence conservation, LL1 and LL2 contrast with one 
another in noteworthy ways. On one hand, the LL1 structure varies considerably in length 
ranging from 37 to76. Across this structure there are stretches of sequence that show little if any 
conservation, but there are also portions which are virtually invariant across all catalase-
peroxidases, such as the ?MGLIYVNPEG? sequence that appears at the LL1 C-terminus (98). In 
contrast, LL2 varies little in its length, and yet at the same time, strictly conserved residues are 
far less prevalent in the structure than in LL1.   
LL1 is positioned at one edge of the heme between D and E helices, and it connects the 
distal and proximal sides of the active site (91-94). The strictly conserved residues, e.g., Tyr249, 
Ile248, and Glu253 (Synechocystis numbering), have been identified as essential for the H2O2 
 
136 
  
oxidation required for catalase turnover (91-94, 102). Tyrosine 249 participates in a unique 
Trp122-Tyr249-Met275 covalent adduct found on the distal side of the active site heme. Site-
specific substitutions preventing formation of the full adduct (e.g., Y249F (90), M275I (114), 
etc.) produce KatG variants with substantially diminished catalase activity. It has been reported 
that Ile248 maintains the heme architecture by forming a hydrogen bond with Asp152 (105), and 
Glu253 creates an acidic entrance to the access channel, which helpd to stabilize and orients the 
solute matrix essential for H2O2 oxidation (102).  
Crystal structures of KatGs indicate that LL2 begins and terminates adjacent to the active 
site between the F and G helices. The ascending and descending limbs of LL2 run roughly 
parallel to a hydrogen bond network formed by distal histidine (His106), Asn136, and Leu130. A 
KatG variant entirely lacking LL2 showed a 200-fold decrease in catalase activity, but only a two 
fold reduction in peroxidase activity (87). However, the mechanism by which LL2 exerts active 
site control is still to be explored as the majority of effort has focused on understanding the role 
of LL1 in the catalytic versatility of KatG.  
  A consistent feature of LL2 in all solved structures for KatG enzymes is the contact of 
its apex with the C-terminal domain of another subunit. The LL2 apex contains six amino acids 
(Ala288-Pro289-Ile290-Glu291-Glu292-Gln293, E.coli residues and numbering) which interact 
with a hydrophobic cavity of C-terminal domain (Figure 4.1). In order to investigate the role of 
this point of contact in maintaining the active site architecture and catalytic ability of E.coli 
KatG, a series of site-specific and deletion variants were generated and the catalytic and 
spectroscopic properties were evaluated.  
 
137 
  
 
 
Figure 4.1: Position of LL2 with respect to the active site Heme. Structure was taken from 
PDB accession domain 1SJ2. LL2 starts and ends adjacent to the active site. IIe290, Glu291, and 
Gln293 (E.coli numbering) are located on the apex of LL2 forming a point of contact with 
hydrophobic cavity of C-terminal domain of another subunit. The residues investigated in this 
study are highlighted in orange.  
 
G 2 9 4
C - term i na l  hy dro pho bi c P o cket
I 2 9 0Q 2 9 3
E2 9 1
P 2 8 9
P 2 8 5
N 2 8 0
S 2 9 9
N 1 3 6
H 1 0 6
H 2 6 7
W3 1 9
D 3 7 7
LL2
LL1
 
138 
  
 
 
 
Table 4.1: Classification of variants based on catalatic activity 
Group I          Group II Group III 
I290A KatG 
Q293A KatG 
I290A/Q293A KatG 
I290V KatG 
Q293V KatG 
I290V/Q293V KatG 
Q293E KatG 
E291A KatG 
E291K KatG 
E291K/E292A KatG 
IEEQ-A KatG 
KatG?291 
 
KatG?P285/?G294 
 
KatG?N280/?S299 
 
KatG?288-293 
 
KatG?290-293 
 
KatG?291-292 
 
KatG?LL2 
 
 
 
139 
  
The variants settled into three groups on the basis of their catalase activity (Table 4.1). 
Most variants generated by site-directed mutagenesis (i.e., substitution variants) showed similar 
catalase activity to wtKatG. Likewise, their spectroscopic characteristics changed little compared 
to wtKatG, suggesting little if any effect on the heme environment.  Consistent with these results, 
the kinetics of peroxidase activity and CN- binding were all similar to one another and wtKatG.  
Conversely, the majority of deletion variants showed catalytic and spectroscopic characteristics 
almost identical to the variant lacking the entire  LL2 structure (87).  The deletion variants had 
clearly impaired catalase activity and an altered active site structure. These proteins were 
dominated by high-spin pentacoordinate heme rather than the typical hexacoordinate state heme-
dominated wtKatG. These results suggest that the strict length conservation of LL2 plays an 
important role in the formation of intersubunit interactions between LL2 apex and C-terminal 
domain; this interaction in turn maintains the appropriate heme environment for the 
multifunctional activity of KatG.  
4.2. Materials and Methods: 
4.2.1.Materials 
All oligonucelotide primers were obtained from Invitrogen (Carlsbad, CA). E.coli strains 
(XL-1 Blue and BL-21 [DE3]) and Pfu polymerase were purchased from Agilent Technologies 
(La Jolla, CA).  Restriction enzymes were obtained from New England Biolabs (Beverly, MA). 
QIAprep Spin Miniprep kit and nickel-nitriloacetic acid (Ni-NTA) resin were obtained from 
Qiagen (Valencia,CA). Isopropyl-B-D-thioglactopyranoside (IPTG) was obtained from Gold 
 
140 
  
Biotechnology Inc. (St. Louis, MO). Mono and di-basic sodium phosphate, acetic acid, and 
sodium acetate were purchased from Fisher (Pittsburgh,PA). Bugbuster and benzonase nuclease 
were purchased from EMD (Madison, WI). Ampicillin, chloramphenicol, imidazole, hemin, 
sodium dithionite, 2,2?-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) (ABTS), and hydrogen 
peroxide (30%), were obtained from Sigma (St. Louis, MO). Water purified through Barnstead 
Easy Pure II UV ultrapure water system (18.2 M?/cm resistivity) was used to make all buffers 
and media. 
4.2.2. Cloning of Site-specific and deletion variants 
The site-specific and deletion variants of the LL2 were generated using the ?around the 
horn? approach as described in chapter 2, with the exception of  I290V KatG and I290V/Q2393V 
KatG. For these two variants the QuikChangeTM procedure was used. Table 4.2 lists the 
mutagenic primers used in this study. Amplification products were used to transform the E. coli 
XL-1 Blue by heat shock procedure. Colonies were selected on the basis of ampicillin resistance, 
and the desired plasmids were subjected to restriction endonuclease digestion as successful 
mutants must contain a specific alteration to the restriction endonuclease digest pattern (Table 
4.2). Each restriction site is an addition either by silent mutation or by specific mutation. 
Restriction digest-positive candidates were subjected to DNA sequencing analysis to confirm 
that the desired mutation had occurred and no undesired mutations had taken place.  
 
 
 
141 
  
Table 4.2: List of primers used for making site-directed and deletion LL2 variants 
Variant Forward Primer Reverse Primer Restriction Enzyme  
I290V KatG 5?-Phos- GCTGCACCGGTTGAAGAACAA-3? 5?-Phos- TTGTTCTTCAACCGGTGCAGC-3?  Age I 
Q293V KatG 5?-Phos- CCGATTGAAGAAGTTGGCCTAGGTTGGGCGAGC-3? 5?-Phos - GCTCGCCCAACCTAGGCCAACTTCTTCAATCGG-3? Sty I 
I290V/Q293V KatG 5?-Phos - GCTGCACCGGTTGAAGAAGTTGGTTTAGGTTGG-3? 5?-Phos - CCAACCTAAACCAACTTCTTCAACCGGTGCAGC-3? Age I 
I290A KatG 5?-Phos -CCGGCGGAAGAACAAGGTTTAGGTTGGGCG-3? 5?-Phos - CGCAGCTTCTGGATCAGGACCTACATTTGATGTC-3? Nae I 
Q293A KatG 5?-Phos - CCGATTGAAGAAGCAGGCCTAGGTTGGGCG-3? 5?-Phos - CGCAGCTTCTGGATCAGGACCTACATTTGATGTC-3? Sty I 
I290A/Q293A KatG 5?-Phos - CCGGCGGAAGAAGCGGGTTTAGGTTGG-3? 5?-Phos - CGCAGCTTCTGGATCAGGACCTACATTTGATGTC-3? Nae I 
E291A KatG 5?-Phos - CCGATCGCGGAACAAGGTTTAGGTTGGGCG-3? 5?-Phos - CGCAGCTTCTGGATCAGGACCTACATTTGATGTC-3? Pvu I 
E291K KatG 5?-Phos- CCGATTAAAGAACAAGGTTTAGGTTGGGCGAGC-3? 5?-Phos - CGCAGCTTCTGGATCAGGACCTACATTTGATGTC-3? N/A 
E291K/E291A KatG 5?-Phos ? CCGATTAAAGCCCAAGGTTTAGGTTGGGCG-3? 5?-Phos - CGCAGCTTCTGGATCAGGACCTACATTTGATGTC-3? Sty I 
Q293E KatG 5?-Phos - CCGATTGAAGAAGAAGGCCTAGGTTGGGCG-3? 5?-Phos - CGCAGCTTCTGGATCAGGACCTACATTTGATGTC-3? Sty I 
KatG?288-293 5?-Phos - GGCTTAGGTTGGGCGAGCACTTACGGC-3? 5?-Phos - GGCTTCTGGATCAGGACCTACATTTGATGTCGG-3? Nae I 
KatG?290-293 5?-Phos - GGTTTAGGTTGGGCGAGCACTTACGGCAGC-3? 5?-Phos - CGGGGCAGCTTCTGGATCAGGACCTAC-3? Sma I 
KatG?291-292 5?-Phos- CCGATCCAAGGTTTAGGTTGGGCGAGC-3? 5?-Phos - CGCAGCTTCTGGATCAGGACCTACATTTGATGTC-3? Sty I 
KatG?291 5?-Phos - CCGATCGAACAAGGTTTAGGTTGGGCGAGC-3? 5?-Phos - CGCAGCTTCTGGATCAGGACCTACATTTGATGTC-3? Pvu I 
KatG?285/?294 5?-Phos - GCGCCGATTGAAGAACAATTGGGTTGGGCGAGC-3? 5?-Phos- CGCTTCATCAGGACCTACATTTGATGTCGGACCG-3? Kas I / Sty I 
IEEQ-A KatG 5?-Phos -  CCGGCGGCTGCAGCGGGTTTAGG-3? 5?-Phos - CGCAGCTTCTGGATCAGGACCTACATTTGATGTC-3? Pst I 
KatG?280/?299 
KatG ?280 
KatG ?299 
 
 
5?-Phos- GGTCCGACGTCAGTAGGTCCTGATCC-3? 
5?-Phos- GCGACGTACGGCAGCGGCGTTGG-3? 
(used  KatG ?280 as template) 
 
5?-Phos- GGCACCGTGGGTTTTACCCAGCGTATG-3? 
5?-Phos- CCAACCTAAACCTTGTTCTTCAATCGGTGC-3? 
 
Aat II 
 
BsiW I 
 
142 
  
4.2.3. Expression and purification 
Expression and purification of all the variants were carried out as described in chapter 2. 
Concentrations of purified enzyme was estimated according to the method of Gill and von 
Hippel (126) using ?280= 1.44 x 105 M-1cm-1  for all the variants with the exception of KatG?LL2, 
where ?280= 1.37 x 105 M-1cm-1  was used.   
4.2.4. Circular Dichroism  
Far-UV CD spectra (180-300 nm) were recorded for all the variants prior to 
reconstitution with hemin by using 5 ?M of each enzyme in 5 mM phosphate buffer, pH 7.0. A 
Jasco J-180 spectropolarimeter was used to record all CD spectra by using 0.05 cm pathlength 
quartz cell, 0.05 nm data pitch, continuous scanning mode, scanning speed of 50 nm/min, and 8 
accumulations. Baseline correction and data analysis were done using J-720 software from Jasco.  
4.2.5. UV-visible absorption spectra and activity assays 
Wild-type KatG and its variants were reconstituted using 0.9 equivalents of hemin.  
Following heme addition, each protein was allowed to incubate for 24 hrs at 4  C. After 24 hrs, 
each reconstituted protein was centrifuged at 13,000 rpm for 5 minutes to remove any free hemin 
or insoluble proteins. All spectra were recorded between 800 nm to 250 nm at room temperature 
in 200 mM phosphate buffer, pH 7.0 using Shmiadzu UV-1601 spectrophotometer. Method of 
Falk was used to determine the concentration of reconstituted enzyme (124). 
 
143 
  
Catalase activity was evaluated by monitoring the decrease in H2O2 concentration at 240 
nm (?240= 39.4 M-1 cm-1) (135). Peroxidase activity was measured by monitoring the production 
of ABTS radical at 417 nm (?417 = 34.7 mM-1 cm-1), initial concentration of ABTS was measured 
spectrophotometrically at 340 nm (?340 = 3.66 x 104 M-1 cm-1) (136). All assays were carried out 
at room temperature using Shimadzu UV-1601 spectrophotometer using 10 nM enzyme, 200 
mM phosphate buffer, pH 7.0. 
4.2.6. Magnetic Circular Dichroism 
A Jasco J-810 spectropolarimeter was used to record MCD spectra. All spectra were 
recorded using 15 ?M of enzyme in 50 mM phosphate buffer, pH 7.0, and 50 mM NaCl. 
Samples were placed in a 0.5 cm pathlength cuvette, and the cuvette was placed in a magnetic 
cell holder with in 1.4 Tesla magnetic field generated by permanent magnet. All spectra were 
recorded at room temperature using the following instrument settings: wavelength, 700 nm to 
350 nm; scanning mode, continuous;  scanning speed, 50 nm/min; data pitch, 0.1 nm; and 
accumulations, 8. In order to obtain MCD spectra for the ferrous heme state, small amounts < 1 
mg solid sodium dithionite was added to the sample cuvette and mixed. Only four spectral 
accumulations were collected for the ferrous state.  The baseline correction and data analysis 
were done using Jasco J-720 software. For baseline correction, buffer spectrum was subtracted 
from the MCD spectrum, and the difference spectrum was used for data analysis. The data were 
fit to equation 4.1. 
?   ?H =            (4.1)      D e l t a  O D           ( E n z y m e  c o n c e n t r a t i o n  m M )  x  1 . 4  T e s l a  x  0 . 5  c m  p a t h l e n g t h
 
144 
  
4.2.7. Electron Paramagnetic Resonance Spectroscopy and Spin Quantification 
EPR spectra were recorded using Bruker EMX instrument equipped with an Oxford ESR 
900 cryostat equipped with an ITC temperature controller. Samples were concentrated to a final 
concentration of 150 ?M in a final volume of 300 ?l using Amicon Ultra-4 centrifuge filter 
devices. Spectrometer settings were as follows: temperature, 10K; center field, 3250 G; sweep 
width, 5500 g; start field, 3250 G; receiver gain, 1.0 x 105; microwave power, 9.38 GHz; 
modulation amplitude, 10 G; modulation frequency, 100 kHz; time constant, 655.36 ms; and 
conversion time, 655.36 ms. Spin quantification was carried out as described in chapter 2 using 
Biomolecular EPR spectroscopy software package available online (134). 
4.2.8. Evaluation of cyanide binding by titration and by stopped-flow 
Cyanide binding was evaluated by two methods: equilibrium titration, and stopped-flow 
kinetics. 
Cyanide binding by titration was monitored using Shimadzu UV-1601 
spectrophotometer. Titrations were performed by adding 1-2 ?L aliquots of a potassium cyanide 
stock solution to 1 mL of each variant (1?M) held in a cuvette. After each portion of cyanide was 
added, the solution was thoroughly mixed, and the optical spectrum was recorded after 3 min to 
ensure completion of the binding reaction. Change in absorbance at 422 nm was plotted against 
[KCN] to obtain the dissociation constants. All cyanide binding studies were carried out in 50 
mM phosphate buffer, pH 7.0, at room temperature.  
 
145 
  
For stopped-flow kinetics, SX.18 MV Stopped-flow Rapid Reaction Analyzer in single 
mixing mode was used. One syringe contained 6 ?M of enzyme and the second contained KCN 
(0.03-3 mM). Spectra were recorded by diode array. Kinetic constants were determined form 
triplicate measurements of single wavelength data recorded at 400 nm (decay of high-spin ferric 
species) and at 428 nm (formation of low-spin hexacoordinate ferric species). All reactions were 
carried out at 25 C in 50 mM phosphate buffer, pH 7.0. 
4.3. Results 
Alignments were performed using sequences of 20 KatGs from across the phylogentic 
spectrum. Two or more representative samples were taken from each group viz., fungal, bacterial 
and archeal groups. The samples were chosen on the basis of their separation in phylogenetic 
relationship tree for catalase-peroxidases provided by Obinger, et.al. (163). The degree to which 
the sequence and length of three large loops are conserved is shown in Figure 4.2. The sequence 
alignment shows that the length of LL1 varied from 37-76 amino-acids among KatGs contained 
~10 highly conserved amino acid residues. In contrast, LL2 was highly conserved in its length 
(35 amino-acids) with very little (4 invariant amino acids) residue conservation. Three out of the 
four strictly conserved residues in LL2, were either proline or glycine. These residues tend to 
have greatest impact on conformation rather than on catalysis. Also, the other strongly conserved 
residues are almost all hydrophobic (i.e. conservation is between otherwise hydrophobically 
equivalent residues viz. Leu, Ile, Val, Met, Phe, etc.). The only other strongly conserved residue 
which did not fit the second category is tryptophan at position 297. We did not observe any 
apparent structural or functional effects on substituting W297 to alanine. The W297A variant 
 
146 
  
behaved the same as wtKatG by having equivalent activity profile and spectroscopic features. In 
contrast, the strictly conserved residues of LL1 included several aromatic residues which have 
shown to be crucial for bifunctionality of KatGs (90, 107, 116). This indicated that length 
conservation of LL2 over residue conservation might be an important criterion in catalytic 
versatility of KatG.  
Previously, Goodwin lab has shown that the LL2 structure is required to maintain the 
bifunctionality of KatG (87, 103). Indeed, the variant lacking the entire LL2 structure caused a 
two orders of magnitude loss of catalase activity but only two-fold reduction in peroxidase 
activity and also diminished the rate of cyanide binding (87). However, the mechanism behind 
this observation was not clearly understood. Closer examination of the LL2 structure indicated 
that the ascending and descending limbs of this loop run roughly parallel to the hydrogen bond 
network formed by distal histidine (His106), Asn136, and Leu130. Thus, removal of entire LL2 
was suggested to disrupt the above mentioned hydrogen bond network as two of its residues 
Glu284 and Trp297 lie fairly close to this network (87, 103). 
Given the essential nature of LL2, its lack of sequence conservation but striking length 
conservation, and the prominent interaction with the C-terminal domain at the apex of the 
structure, we targeted LL2 in two different ways. First, we targeted the individual residues (I290, 
E291, E292, and Q293) which make an intersubunit contact with the hydrophobic cavity of C-
terminal domain. These residues were substituted individually and in combination using site-
directed mutagenesis. The second approach was to target the length of LL2 by deleting residues 
either at its apex (A288-Q293) or along its ascending and descending limbs (P285 and G294,                  
 
147 
  
A. 
 
 
Figure 4.2: Multiple sequence alignment of 20 selected representatives from Class I families with one sequence from a typical 
Class II representative. Presented here are the three large loops of KatG; LL1 (A), LL2 (B), and LL3 (C).  ClustalX version 
2.0.11 was used as a tool for multiple alignments of protein sequences (164, 165). Dotted line represents length of the loops and 
arrows indicate their direction. ?*? represents positions which have single, fully conserved residue, ??? represents that one of the 
?stronger? [STA, NEQK, NHQK, NDEQ, QHRK, MILV, MILF, HY, FYM] group is fully conserved, ?:? represents that one of the 
?weaker? [CSA, ATV, SAG, STNK, STPA, SGND, SNDEQK, NDEQHK, NDEQHRK, FVLIM, HFY] group is fully conserved. 
Abbreviations of species correspond with their names in UniProtKB (166, 167).  
 
148 
  
B.
         
C. 
 
 
 
 
 
 
 
Figure 4.2: (Continued) 
 
149 
  
N280 and S299) (Figure 4.1). Each vectors for the expression of each variant was successfully 
produced and confirmed by DNA sequence analysis. Each of the variants were over-expressed in 
a soluble form and showed proper global folding of the protein as indicated by circular dichroism 
spectra when compared with the wtKatG (Figure 4.3). 
4.3.1. Catalase activity 
The catalase activity of the variants was compared with respect to the wtKatG and the 
variant lacking LL2 (KatG?LL2) (Table 4.3). The variants divided into three distinct groups based 
on their catalatic ability. The apparent second order rate constants (kcat/KM) of Group I variants 
were similar to wtKatG (i.e., 106 M-1s-1) suggesting minimal loss of catalase activity in these 
variants (Figure 4.4). In stark contrast, Group III variants showed catalase activity at least two 
orders of magnitude lower than wtKatG as indicated by their low turnover numbers (i.e., kcat) 
(Figure 4.5). The apparent kcat/KM  for group III variants was three orders of magnitude lower 
than wtKatG, but was similar to the variant which lacked the entire LL2 structure (87). The 
middle group of variants (i.e., Group II) retained minimal catalase activity as 80-90% reduction 
in their apparent kcat values was observed when compared to wtKatG (Figure 4.6).  
4.3.2. Peroxidase activity 
The kinetic parameters for peroxidase activity with respect to H2O2 and ABTS are shown 
in Tables 4.4 and 4.5, respectively. The data obtained from activity assays were fit to standard  
 
 
150 
  
190 210 230 250
- 2 0 0 0 0
- 1 0 0 0 0
0
10000
20000
30000
w t K a t G
I 2 9 0 A  K a t G
Q 2 9 3 A  K a t G
I 2 9 0 A / Q 2 9 3 A  K a t G
E 2 9 1 K  K a t G
A E 2 9 1 A  K a t G
W a v e le n g t h  ( n m )
[
?
]
 
d
e
g
 
c
m
2
d
m
ol
-1
190 210 230 250
- 2 0 0 0 0
- 1 0 0 0 0
0
10000
20000
30000
w t K a t G
I 2 9 0 V  K a t G
Q 2 9 3 V  K a t G
I 2 9 0 V / Q 2 9 3 V  K a t G
Q 2 9 3 E  K a t G
B E 2 9 1 K / E 2 9 2 A  K a t G
W a v eleng t h ( nm )
[
?
]
 
deg
 
cm
2
dm
o
l
-1
190 210 230 250
- 2 0 0 0 0
- 1 0 0 0 0
0
10000
20000
30000
w t K a t G
K a t G
? P 2 8 5 / ? G 2 9 4
K a t G
? 291
K a t G
? N2 8 0 / ? S 2 9 9
I E E Q- A  K a t G
C
W a v e le ng t h ( nm)
[
?
]
 
de
g
 
c
m
2
d
m
o
l
-1
190 210 230 250
- 2 0 0 0 0
- 1 0 0 0 0
0
10000
20000
30000
w t K a t G
K a t G
? LL2
K a t G
? 2 8 8 - 2 9 3
K a t G
? 2 9 0 - 2 9 3
K a t G
? 2 9 1 - 2 9 2
D
W a v e le ng t h ( nm )
[
?
]
 
de
g
 
c
m
2
d
m
ol
-1
 
Figure 4.3: CD spectra of group I (A and B), group II (C), and group III (D) variants. Each spectrum was recorded with apo- 
enzyme in 5 mM phosphate buffer, pH 7.0 at room temperature. 
 
151 
  
Table 4.3: Apparent catalase kinetic parameters 
 
 
 
KatG 
 
Catalase cycle parameters  
kcat(s-1) KM (mM H2O2) kcat/KM (106 M-1s-1) Fraction (kcat) 
  WT KatG 11000 ? 200 3.5 ? 0.2 3.2 1 
  I290A KatG 10173 ? 309 3.7 ? 0.3 2.7 0.92 
  Q293A KatG 12176 ? 621 4.6 ? 0.6 2.6 1.10 
  I290A/Q293A KatG 7606 ? 440 4.1 ? 0.7 1.9 0.69 
Group I I290V KatG 
 
7981 ? 187 3.9 ? 0.3 2 0.72 
  Q293V KatG 
 
5309 ? 233 3.1 ? 0.4 1.7 0.48 
  I290V/Q293V KatG 9540 ? 293 3.2 ? 0.4 1.8 0.86 
  Q293E KatG 
 
9624 ? 568 8 ? 1 1.2 0.87 
  E291A KatG 16653 ? 487 5.9 ? 0.4 2.8 1.51 
  E291K KatG 
 
9540 ? 293 2.1 ? 0.2 4.5 0.86 
  E291A/E2912K KatG 12184 ? 390 2.5 ? 0.3 4.8 1.10 
  IEEQ-A KatG 
 
626 ? 28 3.4 ? 0.5 0.18 0.05 
  KatG?291 
 
1983 ? 47 3.1 ? 0.2 0.64 0.18 
Group II KatG?P285/?G294 1766 ? 49 2.2 ? 0.2 0.8 0.16 
  KatG?N280/?S299 
 
2535 ? 114 3.2 ? 0.4 0.79 0.23 
  KatG?288-293 
 
7 ? 7 3 ? 14 0.0023 0.0006 
Group III KatG?290-293 37 ? 5 5 ? 1 0.0074 0.0033 
 
  KatG?291-292 
 
KatG?LL2 
 
N/D 
 
N/D 
N/D 
 
N/D 
N/D 
 
N/D 
N/D 
 
N/D 
N/D: Not Detected  
 
152 
  
Michaelis-Menten equation (4.1). However, in some variants substrate-dependent inhibition was 
observed and the data were fit to equation 4.2. 
v = k c a t  [ S ]
K M  +  [ S ][ E ] T
( 4 . 1 )
 
=
k c a t  [ S ]
K M  +  [ S ]  +  [ S ] 2   K I
v
[ E ] T
( 4 . 2 )
 
   In case of peroxidase activity with respect to H2O2 concentration, a two to four- fold 
reduction in apparent kcat was observed for group I variants with a much more dramatic decrease 
in apparent KM (Figure 4.7, Panel A).  This decrease in both apparent kcat and apparent KM gave 
rise to similar apparent kcat/KM values compared to wtKatG. Conversely, group II (Figure 4.8, 
Panel A) and III (Figure 4.9, Panel A) variants showed a two to four-fold higher apparent kcat 
with much higher apparent KM values. The net result with respect to kcat/KM was 10-fold lower 
apparent second-order rate constants for these variants.  
The apparent kcat values measured for group I variants for ABTS concentration were 
reduced by two to four-fold with modest decrease in apparent KM except some cases where it was 
either equal to (Q293A, Q293V, and I290V/Q293V) or slightly higher (I290A and I290V) than 
that for wtKatG (Figure 4.7, Panel B). In case of the group II (Figure 4.8, Panel B)  and III 
(Figure 4.9, Panel B) variants, there was slight increase in apparent kcat and decrease in KM 
leading to overall higher apparent second order rate constant (~106 M-1s-1), with the exception of 
KatG?P285/?G294 and KatG?N280/?S299 which showed ~40 % decrease in apparent kcat/KM.  
 
153 
  
 
 
 
 
Figure 4.4: Effect of H2O2 concentration on the catalase activity of wtKatG and group I 
variants. Reactions were carried our using 10 nM enzyme and 0.2-20 mM H2O2 in 100 mM 
phosphate buffer, pH 7.0, at 23?C 
 
 
 
 
0 5 10 15 20
0
2000
4000
6000
8000
10000
12000
14000
I 2 9 0 A  K a t G
Q 2 9 3 A  K a t G
I 2 9 0 A / Q 2 9 3 A K a t G
I 2 9 0 V  K a t G
Q 2 9 3 V  K a t G
I 2 9 0 V / Q 2 9 3 V  K a t G
Q 2 9 3 E  K a t G
E 2 9 1 A  K a t G
E 2 9 1 K  K a t G
E 2 9 1 K / E 2 9 2 A  K a t G
w t K a t G
[H
2
O
2
] ( m M )
v
o
/[
E
]
T
(s
-1
)
 
154 
  
 
 
 
 
0 5 10 15 20
- 1 0 0 0
1000
3000
5000
7000
9000
11000
13000
K a t G
? LL2
K a t G
? 2 8 8 - 2 9 3
K a t G
? 2 9 0 - 2 9 3
K a t G
? 2 9 1 - 2 9 2
w t K a t G
[H
2
O
2
] ( m M )
v
o
/[
E
]
T
(s
-1
)
 
 
Figure 4.5: Effect of H2O2 concentration on the catalase activity of wtKatG, KatG?LL2 and 
group III variants. Reactions were carried our using 10 nM enzyme and 0.2-20 mM H2O2 in 
100 mM phosphate buffer, pH 7.0, at 23?C. 
 
 
 
 
155 
  
 
 
 
0 5 10 15 20
0
2000
4000
6000
8000
10000
12000
I E E Q - A  K a t G
K a t G
? 291
K a t G
? N2 8 0 / ? S 2 9 9
w t K a t G
K a t G
? P 2 8 5 / ? G 2 9 4
[H
2
O
2
]  ( m M )
v
o
/[
E
]
T
(s
-1
)
 
Figure 4.6: Effect of H2O2 concentration on the catalase activity of wtKatG and group II 
variants. Reactions were carried our using 10 nM enzyme and 0.2-20 mM H2O2 in 100 mM 
phosphate buffer, pH 7.0, at 23?C. 
 
 
 
 
 
 
 
156 
  
Table 4.4: Apparent peroxide-dependent peroxidase kinetic parameters 
KatG Peroxide-dependent peroxidase cycle parameters  
kcat(s-1) KM (mM H2O2) kcat/KM (105 M-1s-1) KI(mM H2O2) 
 
 
 
 
Group I 
WT KatG 
I290A KatG 
Q293A KatG 
I290A/Q293A KatG 
I290V KatG 
Q293V KatG 
I290V/Q293V KatG 
Q293E KatG 
E291A KatG 
E291K KatG 
E291A/E292K KatG 
65 ? 6 
26.3 ? 0.6 
27.8 ? 0.6 
13.5 ? 0.4 
14.66 ? 0.3 
22.1 ?  1.3 
22 ? 1 
23.8 ? 0.5 
40.4 ? 0.9 
34.1 ? 0.8 
35.3 ? 0.5 
0.18 ? 0.04 
0.08 ? 0.01 
0.07 ? 0.01 
0.07 ? 0.01 
0.03 ? 0.01 
0.30 ? 0.06 
0.14 ? 0.03 
0.089 ? 0.008 
0.104 ? 0.007 
0.06 ? 0.01 
0.057 ? 0.004 
3.6 
3.3 
3.9 
1.9 
4.9 
0.73 
1.6 
2.7 
3.9 
5.7 
5.9 
N/D 
N/D 
N/D 
N/D 
N/D 
N/D 
N/D 
N/D 
12 ? 1 
N/D 
N/D 
 
 
Group II 
IEEQ-A KatG 
KatG?291 
KatG?P285/?G294 
KatG?N280/?S299 
97 ? 6 
184 ? 8 
247 ? 44 
218 ? 9 
73 ? 9 
40 ? 3 
139 ? 32 
56 ? 8 
0.013 
0.046 
0.018 
0.039 
N/D 
N/D 
N/D 
N/D 
 
Group III 
KatG?288-293 
KatG?290-293 
KatG?291-292 
KatG?LL2 
46 ? 1.8 
422 ? 107 
261 ? 19 
27.8 ? 0.2 
7.7 ? 0.7 
215 ? 65 
98 ? 12 
2.2? 0.2 
0.059 
0.019 
0.027 
0.13 
N/D 
N/D 
N/D 
N/D 
N/D: Not Detected  
 
 
157 
  
Table 4.5: Apparent ABTS-dependent peroxidase kinetic parameters 
 
KatG 
ABTS-dependent peroxidase cycle parameters  
kcat(s-1) KM (mM ABTS) kcat/KM (105 M-1s-1) KI (mM ABTS) 
 
 
 
 
Group I 
WT KatG 
I290A KatG 
Q293A KatG 
I290A/Q293A KatG 
I290V KatG 
Q293V KatG 
I290V/Q293V KatG 
Q293E KatG 
E291A KatG 
E291K KatG 
E291K/E292A KatG 
55.2 ? 1.3 
32.5 ? 0.9 
29.7 ? 0.5 
13.3 ? 0.4 
28.4 ? 0.7 
17.4 ?0.5 
22.7 ? 0.3 
20.2 ? 0.6 
39.7 ? 0.8 
29.6 ? 0.3 
38.2 ? 0.8 
0.087 ? 0.008 
0.16 ? 0.02 
0.08 ? 0.01 
0.037 ? 0.06 
0.13 ? 0.01 
0.09 ? 0.01 
0.09 ? 0.01 
0.052 ? 0.008 
0.07 ? 0.01 
0.023 ? 0.02 
0.030?0.003 
6.3 
2.0 
3.7 
3.6 
2.2 
1.9 
2.7 
3.9 
5.7 
13 
13 
N/D 
N/D 
N/D 
N/D 
N/D 
N/D 
N/D 
N/D 
N/D 
N/D 
N/D 
 
 
Group II 
IEEQ-A KatG 
KatG?291 
KatG?P285/?G294 
KatG?N280/?S299 
70 ? 3 
68 ? 1 
29 ? 1 
54.3 ? 0.9 
0.029 ? 0.003 
0.062 ? 0.05 
0.069 ? 0.007 
0.074 ? 0.005 
24 
11 
4.2 
7.3 
0.9 ? 0.2 
N/D 
4.3 ? 0.8 
N/D 
 
Group III 
KatG?288-293 
KatG?290-293 
KatG?291-292 
KatG?LL2 
65 ? 1 
37.7 ? 0.9 
54 ? 1 
25.7 ? 0.9 
0.041 ? 0.004 
0.022 ? 0.002 
0.019 ? 0.002 
0.012 ? 0.001 
16 
17 
28 
22 
N/D 
N/D 
N/D 
N/D 
N/D: Not Detected 
   
 
158 
  
0 . 0 0 . 5 1 . 0 1 . 5 2 . 0
0
5
10
15
20
25
30
35
40
45
I 2 9 0 A  K a t G
Q 2 9 3 A  K a t G
I 2 9 0 A / Q 2 9 3 A  K a t G
I 2 9 0 V  K a t G
Q 2 9 3 V  K a t G
I 2 9 0 V / Q 2 9 3 V  K a t G
Q 2 9 3 E  K a t G
E 2 9 1 A  K a t G
E 2 9 1 K  K a t G
E 2 9 1 K / E 2 9 A  K a t G
w t K a t G
[H
2
O
2
]  ( m M )
v
o
/[
E
]
T
(s
-1
)
A
0 . 0 0 . 5 1 . 0 1 . 5 2 . 0
0
10
20
30
40
50
I 2 9 0 A  K a t G
Q 2 9 3 A  K a t G
I 2 9 0 A / Q 2 9 3 A  K a t G
I 2 9 0 V  K a t G
Q 2 9 3 V  K a t G
I 2 9 0 V / Q 2 9 3 V  K a t G
Q 2 9 3 E  K a t G
E 2 9 1 A  K a t G
E 2 9 1 K  K a t G
E 2 9 1 K / E 2 9 2 A  K a t G
w t K a t G
B
[ AB T S ]  ( m M )
v
o
/[
E
]
T
(s
-1
)
 
Figure 4.7: Comparison of peroxidase activities of wtKatG and group I variants. The effects 
of H2O2 (A) and ABTS (B) concentrations on the rate of ABTS oxidation by wtKatG and group 
III variants were evaluated. (A) All reactions contained 0.5 mM ABTS. (B) All reactions 
contained 0.5 mM H2O2. All reactions were carried out at 23?C in 50 mM acetate buffer, pH 5.0, 
using 10 nM of enzyme.  
 
159 
  
0 10 20 30 40 50
0
15
30
45
60
75
90
105
I E E Q - A  K a t G
K a t G
? 291
K a t G
? N2 8 0 / ? S 2 9 9
w t K a t G
K a t G
? P 2 8 5 / ? G 2 9 4
[H
2
O
2
]  ( m M )
v
o
/[
E
]
T
(s
-1
)
A
0 .0 0 .5 1 .0 1 .5 2 .0
0
15
30
45
60
75
I E E Q - A  K a t G
K a t G
? 291
K a t G
? N2 8 0 / ? S 2 9 9
w t K a t G
K a t G
? P 2 8 5 / ? G 2 9 4
[ A B T S ]  ( m M )
v
o
/[
E
]
T
(s
-1
)
B
 
Figure 4.8: Comparison of peroxidase activities of wtKatG and group II variants. The 
effects of H2O2 (A) and ABTS (B) concentrations on the rate of ABTS oxidation by wtKatG and 
group III variants were evaluated. (A) All reactions contained 0.5 mM ABTS. (B) 0.5 mM H2O2  
for wtKatG and 20 mM H2O2 for the variants. All reactions were carried out at 23?C in 50 mM 
acetate buffer, pH 5.0, using 10 nM of enzyme.  
 
160 
  
0 10 20 30 40 50
0
20
40
60
80
100
K a t G
? LL2
K a t G
? 2 8 8 - 2 9 3
K a t G
? 2 9 0 - 2 9 3
K a t G
? 2 9 1 - 2 9 2
w t K a t G
[H
2
O
2
]  ( m M )
v
o
/[
E
]
T
(s
-1
)
0 . 0 0 . 5 1 . 0 1 . 5 2 . 0
0
15
30
45
60
75
K a t G
? LL2
K a t G
? 2 8 8 - 2 9 3
K a t G
? 2 9 0 - 2 9 3
K a t G
? 2 9 1 - 2 9 2
w t K a t G
[ AB T S]  ( m M )
v
o
/[
E
]
T
(s
-1
)
 
Figure 4.9: Comparison of peroxidase activities of wtKatG and group III variants. The 
effects of H2O2 (A) and ABTS (B) concentrations on the rate of ABTS oxidation by wtKatG and 
group III variants were evaluated. (A) All reactions contain 0.5 mM ABTS. (B) 0.5 mM H2O2  
for wtKatG and 20 mM H2O2 for the variants. All reactions were carried out at 23?C in 50 mM 
acetate buffer, pH 5.0, using 10 nM of enzyme. 
 
161 
  
4.3.3 Spectroscopic evaluation of KatG LL2 variants 
   The typical UV-Vis spectrum for ferric KatG shows a Soret band absorption maximum at 
408 nm with charge transfer transitions near 500 nm (shoulder) and 642 nm. These features 
generally refer to a mixture of coordination states dominated by hexacoordinate high-spin heme 
with contributions from the pentacoordinate high-spin state (87, 88, 101). Generally, the 
absorption spectra for the ferric forms of the group I substitution variants were highly similar to 
wtKatG (Figure 4.10). For variants in group II (Figure 4.11, Panels A and B) and group III 
(Figure 4.11, Panels C and D), the tendency was for a more substantial shoulder (~380 nm) to the 
Soret band. These variants tended to show a slight red-shift in the charge transfer and CT1 to 645 
nm. These spectral shifts are consistent with a greater contribution from pentacoordinate heme 
states (102, 168). However, similar spectral shifts would be observed for free heme, thus, on the 
basis of UV-Vis absorption spectra alone assignment of coordination environment for these 
variants could not be made.  
Magnetic Circular Dichroism is a useful approach for hemoproteins and because the 
transitions measured are ?- ?? transitions from the porphyrin, it is an approach that can be applied 
at room temperature. Moreover, there is less ambiguity in the interpretation of some aspects of 
the spectra (particularly when applied to the ferrous state) aiding in a more definitive assignment 
of heme coordination. MCD spectra of the ferric state of the variants were consistent with 
domination by high-spin species (Figures 4.12 and 4.13) (168, 169). Each variant was reduced 
with dithionite and MCD spectra for the ferrous state was measured. For the variants in group I, 
the spectra for the ferrous state was virtually invariant. The maximum corresponding to the Soret  
 
162 
  
325 350 375 400 425 450 475
w t K a t G
I 2 9 0 A  K a t G
Q 2 9 3 A  K a t G
E 2 9 1 A  K a t G
E 2 9 1 K  K a t G
E 2 9 1 K / E 2 9 2 A  K a t G
408
380
A
W a v e l e ng t h ( nm )
?
 
(
m
M
-1
cm
-1
)
475 500 525 550 575 600 625 650 675 700
w t K a t G
I 2 9 0 A  K a t G
Q 2 9 3 A  K a t G
E 2 9 1 A  K a t G
E 2 9 1 K  K a t G
E 2 9 1 K / E 2 9 2 A  K a t G
508
642
B
W a v e l e ng t h ( nm )
?
 
(
m
M
-1
cm
-1
)
325 350 375 400 425 450 475
w t K a t G
I 2 9 0 V  K a t G
Q 2 9 3 V  K a t G
I 2 9 0 V / Q 2 9 3 V  K a t G
Q 2 9 3 E  K a t G
I 2 9 0 A / Q 2 9 3 A K a t G
C
408
380
W a v e l e ng t h ( nm )
?
 
(
m
M
-1
cm
-1
)
475 500 525 550 575 600 625 650 675 700
w t K a t G
I 2 9 0 V  K a t G
Q 2 9 3 V  K a t G
I 2 9 0 V / Q 2 9 3 V  K a t G
Q 2 9 3 E  K a t G
I 2 9 0 A / Q 2 9 3 A K a t G
508
642
D
W a v e l e ng t h ( nm )
?
 
(
m
M
-1
cm
-1
)
 
Figure 4.10: UV-Visible spectral comparison of ferric wtKatG and group I variants. Spectra were obtained for ferric wtKatG and 
the variants. Panels (A) and (C) show these spectra from 325 to 475 nm, and Panels (B) and (D) show them from 475 to 700 nm. All 
the spectra were recorded at 23  C in 100 mM phosphate buffer, pH 7.0.  
 
163 
  
325 350 375 400 425 450 475
K a t G
? 291
K a t G
? N2 8 0 / ? S 2 9 9
K a t G
? P 2 8 5 / ? G 2 9 4
I E E Q- A   K a t G
w t  K a t G
408
380
A
W a v e le ng t h ( nm )
?
 
(
m
M
-1
cm
-1
)
475 500 525 550 575 600 625 650 675 700
K a t G
? 291
K a t G
? N2 8 0 / ? S 2 9 9
K a t G
? P 2 8 5 / ? G 2 9 4
I E E Q- A  K a t G
w t  K a t G
508
6 4 2      6 4 5
B
W a v e le ng t h ( nm )
?
 
(
m
M
-1
cm
-1
)
325 350 375 400 425 450 475
K a t G
? 2 8 8 - 2 9 3
K a t G
? 2 9 0 - 2 9 3
K a t G
? 2 9 1 - 2 9 2
K a t G
? LL2
w t  K a t G
408
380
C
W a v e l e n g t h  ( n m)
?
 
(
mM
-1
cm
-1
)
475 500 525 550 575 600 625 650 675 700
K a t G
? 2 8 8 - 2 9 3
K a t G
? 2 9 0 - 2 9 3
K a t G
? 2 9 1 - 2 9 2
K a t G
? LL2
w t  K a t G
508
6 4 2      6 4 5
D
W a v e le ng t h ( nm )
?
 
(
m
M
-1
cm
-1
)
 
Figure 4.11: UV-Visible spectral comparison of ferric wtKatG with group II (A and B), and group III (C and D) variants. 
Spectra were obtained for ferric wtKatG and the variants. Panels (A) and (C) show these spectra from 325 to 475 nm, and Panels (B) 
and (D) show them from 475 to 700 nm. All the spectra were recorded at 25  C in 100 mM phosphate buffer, pH 7.0.
 
164 
  
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
Figure 4.12: Overlay of ferric MCD of wtKatG and group I variants. Spectra were recorded 
at room temperature using 15uM of each enzyme, in 50 mM phosphate buffer, pH 7.0. 
 
350 400 450 500 550 600 650 700
Q 2 9 3 E K a t G
I 2 9 0 V / Q 2 9 3 V  K a t G
Q 2 9 3 V  K a t G
I 2 9 0 V  K a t G
w t K a t G
I 2 9 0 A / Q 2 9 3 A  K a t G
W a v el eng t h ( nm )
??
/
H
 
(
M
-1
cm
-1
 T
-1
)
350 400 450 500 550 600 650 700
I 2 9 0 A  K a t G
Q 2 9 3 A  K a t G
E 2 9 1 A  K a t G
E 2 9 1 K  K a t G
E 2 9 1 A / E 2 9 2 K  K a t G
w t K a t G
W a v e le n g t h  ( n m )
??
/
H
 
(
M
-1
cm
-1
 T
-1
)
 
165 
  
 
 
 
 
 
 
 
 
Figure 4.13: Overlay of ferric MCD wtKatG, group II (A) and group III (B) variants. 
Spectra were recorded at 25  C using 15uM of each enzyme, in 50 mM phosphate buffer, pH 7.0. 
350 400 450 500 550 600 650 700
K a t G
? P 2 8 5 / ? G 2 9 4
I E E Q- A  K a t G
K a t G
? N2 8 0 / ? S 2 9 9
K a t G
? 291
w t K a t G
W a v e leng t h ( nm )
??
/
H
 (
M
-1
cm
-1
T
-1
)
A
350 400 450 500 550 600 650 700
w t K a t G
K a t G
? LL1
K a t G
? 2 8 8 - 2 9 3
K a t G
? 2 9 0 - 2 9 3
K a t G
? 2 9 1 - 2 9 2
B
W a v e leng t h  ( n m )
??
/
H
 (
M
-1
cm
-1
 T
-1
)
 
166 
  
band was singular and consistently observed near 440 nm. Likewise, transitions corresponding to 
the ? and ? bands were consistently observed near at 557 nm and 585 nm (Figure 4.14).  In 
contrast, group II (Figure 4.15) and III (Figure 4.16) were distinct from group I in that the feature 
around 440 nm was split, with a higher intensity feature at ~ 430 nm and a lower intensity 
feature at ~ 445 nm. 
Figure 4.16 represents an overlay of the typical ferrous MCD spectrum of group III 
variants which clearly shows the splitting of Soret band. Interestingly, KatG?290-293 did not show 
the same split feature, but the wavelength maximum was not consistent with wtKatG in that it 
had blue shifted to 432 nm and at 445 nm there was a notable negative cotton effect. Although 
every group II and III variants were distinct form wtKatG, they all (with the exception of 
KatG?290-293) were highly similar to KatG?LL2. 
The coordination states of the three groups were further evaluated by electron 
paramagnetic resonance (EPR) spectroscopy. The EPR spectra of all the variants, except I290A 
KatG, showed mixed populations of high spin ferric heme species with contribution from an 
axial component (g? = 6.00; g? = 1.99) as well as rhombic component (g = 6.64, 4.95, 1.95). 
However, there was a marked difference in distribution of high spin species among the variants. 
Group I variants showed greater contribution from the axial component and comparatively less 
from rhombic component consistent with wtKatG, indicating domination by the hexacoordinate 
high-spin ferric state (Figure 4.17) (101).  In contrast, spectra for the group II (Figure 4.18) and 
III (Figure 4.19) variants were dominated by the rhombic component with a comparatively 
diminished axial component. These spectra bear a striking resemblance to KatG?LL2.  
 
167 
  
370 420 470 520 570 620 670
W a v e l e ng t h ( nm)
??
/
H
 
(
M
-1
cm
-1
 T
-1
)
I 2 9 0 A  K a t G
Q2 9 3 A  K a t G
E 2 9 1 A / E 2 9 2 K  K a t G
E 2 9 1 K  K a t G
E 2 9 1 A  K a t G
w t K a t G
370 420 470 520 570 620 670
W a v el eng t h ( nm )
??
/
H
 
(
M
-1
cm
-1
 T
-1
)
I 2 9 0 A / Q2 9 3 A  K a t G
Q2 9 3 E  K a t G
I 2 9 0 V / Q2 9 3 V  K a t G
Q2 9 3 V  K a t G
I 2 9 0 V  K a t G
w t K a t G
Figure 4.14: Overlay of ferrous MCD spectra wtKatG and group I variants. Spectra 
were recorded at 25  C using 15 ?M of each enzyme, in 50 mM phosphate buffer, pH 7.0. 
 
168 
  
400 410 420 430 440 450 460 470
W a v e le n g t h  ( n m )
??
/
H
 
(
M
-1
cm
-1
 T
-1
)
432
445
434
440
w t K a t G
I E E Q - A  K a t G
K a t G
? 291
K a t G
? P 2 8 5 / ? G 2 9 4
K a t G
? N 2 8 0 / ? S 2 9 9
560 570 580 590 600 610 620
W a v e le n g t h  ( n m )
??
/
H
 
(
M
-1
cm
-1
 T
-1
)
w t K a t G
I E E Q - A  K a t G
K a t G
? 291
K a t G
? P 2 8 5 / ? G 2 9 4
K a t G
? N 2 8 0 / ? S 2 9 9
 
Figure 4.15: Overlay of ferrous MCD comparison of wtKatG, and group II variants. 
Spectra were recorded at 25  C using 15 ?M of each enzyme, in 50 mM phosphate buffer, pH 
7.0. 
 
 
169 
  
400 410 420 430 440 450 460 470
K a t G
? 2 8 8 - 2 9 3
K a t G
? 2 9 0 - 2 9 3
w t K a t G
K a t G
? 2 9 1 - 2 9 2
K a t G
? LL2
440
432
445
W a v e leng t h  ( n m )
??
/H
 (
c
m
-1
M
-1
T
-1
)
560 570 580 590 600 610 620
K a t G
? 2 8 8 - 2 9 3
K a t G
? 2 9 0 - 2 9 3
w t K a t G
K a t G
? 2 9 1 - 2 9 2
K a t G
? LL2
W a v e leng t h  ( n m )
??
/H
 (
c
m
-1
M
-1
T
-1
)
 
Figure 4.16: Overlay of ferrous MCD spectra of wtKatG, KatG?LL2 and group III variants. 
Spectra were recorded at room temperature using 15 ?M of each enzyme, in 50 mM phosphate 
buffer, pH 7.0. 
 
 
170 
  
 
 
Figure 4.17: Comparison of EPR spectra of wtKatG and group I variants. Relative intensity 
magnified by 4 at field strength greater than 1700 Gauss. EPR settings were identical to those 
described in materials and methods. 
 
500 1500 2500 3500 4500 5500
4X
6 . 6
6 . 0
1 . 9 9
1 . 9 6
w t K a t G
Q 2 9 3 E  K a t G
I 2 9 0 A / Q 2 9 3 A  K a t G
I 2 9 0 V / Q 2 9 3 V  K a t G
Q 2 9 3 V  K a t G
I 2 9 0 V  K a t G
F ield  ( G a u s s )
 
171 
  
 
Figure 4.17: Continued 
 
 
500 1500 2500 3500 4500 5500
4X
6 . 6
6 . 0
1 . 9 9
1 . 9 6
w t K a t G
I 2 9 0 A  K a t G
E 2 9 1 A  K a t G
E 2 9 1 A / E 2 9 2 K  K a t G
E 2 9 1 K  K a t G
Q2 9 3 A  K a t G
F ield  ( G a u s s )
 
172 
  
 
Figure 4.18: Comparison of EPR spectra of wtKatG and group II variants. Relative 
intensity magnified by 4 at field strength greater than 1700 Gauss. EPR settings were identical to 
those described in materials and methods. 
500 1500 2500 3500 4500 5500
4X
6 . 6 3
5
. 8
8
4
.
9
7
1 . 9 9
1 . 9 7
F i el d  ( G a u s s )
I EEQ - A  K a t G
K a t G
? N 2 8 0 / ? S 2 9 9
K a t G
? P 2 8 5 / ? G 2 9 4
K a t G
? 291
w t K a t G
 
173 
  
 
Figure 4.19: Comparison of EPR spectra of wtKatG and group III variants. Relative 
intensity magnified by 4 at field strength greater than 1700 Gauss. EPR settings were identical to 
those described in materials and methods. 
500 1500 2500 3500 4500 5500
4X
6 . 6 3
5 . 8 8
4 . 9 7
1 . 9 9
1 . 9 6
F ield  ( G a u s s )
K a t G
? 2 9 1 - 2 9 2
K a t G
? 2 9 0 - 2 9 3
K a t G
? 2 8 8 - 2 9 3
K a t G
? LL2
w t K a t G
 
174 
  
Together, the UV-Vis, MCD, and EPR spectra indicated that substitution variants to LL2 
has little effect on heme coordination, but deletions to the structure substantially shifted 
coordination from hexacoordinate to pentacoordinate high-spin heme. 
 The next step was to quantify the relative proportion of each coordinating species, and in 
order to accomplish this, EPR spectral simulations and integrations was carried out. The most 
accurate simulations were obtained by accounting for three high spin signals and one low-spin 
signal. The high spin signals included: two axial components (AHS1 with g-values of 5.85 and 
1.99, and AHS2 with g-values of 5.69 and 1.99) and one rhombic component (RHS) with g-
values of 6.65, 4.95, and 1.95. The low-spin component was simulated rhombic (RLS) with g-
values of 2.93, 2.27, and 1.53. Table 4.6 represents the ratios of various EPR signals expressed 
as a percentage contribution of each species in wtKatG and each variant. EPR spectra of all the 
variants irrespective of their groups were dominated by high-spin species. The group I variants 
all showed higher percentage contribution from AHS signal over RHS, a feature common with 
wtKatG. I290A/Q293A KatG and the quadruple alanine (IEEQ-A KatG) variants were the 
exceptions with slightly higher RHS state over AHS state. In contrast, all of the deletion variants 
were dominated by the RHS state. In case of KatG?290-293, the RHS:AHS ratio was almost 50:50 
which might have some relation to  no Soret splitting in MCD ferrous spectrum of this variant. 
But the overall higher rhombic contribution in the deletion variants corroborated well with the 
prominent shoulder at 380 nm in the UV-Vis ferric spectra of these variants.  
 
 
 
175 
  
 
Table 4.6: Ratios, expressed as percentages, of various EPR signal observed in wtKatG and 
variants 
 
        
KatG 
  HS - LS RHS - AHS AHS1 - 
AHS2 
AHS1 ? AHS2 ? RHS - 
RLS 
 
Wt KatG 99.98 ? 0.02 14.9 ? 85.9 49.9 ? 35.1 49.9 ? 35.1-14.9 ? 0.02 
 
I290V KatG 99.93 ? 0.07 39.7 ? 60.2 23.4 ? 36.8 23.4 ? 36.8 - 39.7 ? 0.07 
 
Q293V KatG 99.98 ? 0.02 40.3 ? 59.6 22.5 ? 37.0 22.5 ? 37.0 - 40.3 -0.02 
 
I290V/Q293V KatG 97.88 ? 0.07 39.4 ? 58.5 22.6 ? 35.9 22.6 ? 35.9 ? 39.4 ? 0.07 
 
I290A KatG 100 ? 0 11 ? 89 62.3 ? 26.5 62.3 ? 26.5 ? 11 ? 0 
Group I Q293A KatG 99.7 ? 0.3 35.4 ? 64.3 33.8 ? 30.5 33.8 ? 30.5 ? 35.4- 0.3 
 
I290A/Q293A KatG 99.9 ? 0.1 61.3 ? 38.6 19.6 ? 19.0 19.6 ? 19.0 ? 61.3 ? 0.1 
 
Q293E KatG 99.9 ? 0.1 35.1 ? 64.8 36.9 ? 27.8 36.9 ? 27.8 ? 35.1- 0.1 
 
E291A KatG 99.92 ? 0.07 39.1 ? 60.9 24.6 ? 36.3 24.6 ? 36.3 ? 39.1 ? 0.07 
 
E291K KatG 99.8 ? 0.2 26.4 ? 73.4 47.1 ? 26.3 47.1 ? 26.3 ? 26.4 ? 0.2 
 
E291KE292A KatG 99.9 ? 0.1 37.7 ? 62.1 34.4 ? 27.7 34.4 ? 27.7 ? 37.7 ? 0.1 
 IEEQ-A KatG 99.7 ? 0.3 58.2 ? 41.5 21.0 ? 20.4 21.0 ? 20.4 ? 58.2 ? 0.3 
Group II KatG?291 99.7 ? 0.3 54.4 ? 45.3 23.3 ? 21.9 23.3 ? 21.9 ? 54.4 ? 0.3 
 
KatG?P285/?G294 99.8 ? 0.2 58.5 ? 41.2 20.9 ? 20.2 20.9 ? 20.2-58.5 ? 0.2 
 KatG?N280/?S299 99.8 ? 0.2 68.4 ? 31.3 16.2 ? 15.2 16.2 - 15.2 ? 68.4 ? 0.2 
  KatG?LL2 99.9 ? 0.1 77.1 ? 22.9 11.6 ? 11.2 11.6 ? 11.2 ? 77.1 ? 0.1 
 
KatG?288-293 99.7 ? 0.3 73.5 ? 26.2 10.1 ? 16.0 10.1 ? 16.0 ? 73.5 ? 0.3 
GroupIII KatG?290-293 99.3 ? 0.7 56.2 ? 43.0 20.5 ? 22.5  20.5 ? 22.5 ? 56.2 ? 0.7 
  KatG?291-292 99.7 ? 0.3 75.7 ? 24.0 12.0 ? 11.9 12.0 ? 11.9 ? 75.7 ? 0.3 
 
 
176 
  
4.3.4. Cyanide binding studies 
4.3.4.1 Cyanide titrations  
   Cyanide binding is a useful probe to investigate the changes in structure and accessibility 
of the active site. The spectral changes upon the addition of cyanide to wtKatG and variants from 
all the three groups showed a shift in Soret band maximum to 424 nm from 408 nm along with 
appearance of a new band at 540 nm and disappearance of charge transfer bands at 510 and 645 
nm. These features indicated formation of a hexacoordinate low spin FeIII-CN complex. In 
wtKatG and group I variants first evidence of formation of FeIII-CN complex was observed at 
lower concentration of KCN (~ 2 ?M) (Figure 4.20, A and B). In contrast, the variants from 
group II and III showed an initial decrease in absorbance at 380 nm at lower KCN concentration 
followed by a shift in Soret absorption maximum to 424 nm at higher KCN concentrations (> 10 
?M) (Figure 4.20, C and D). The shift of Soret absorption maximum was accompanied by a 
small decrease in intensity in wtKatG and group I variants, whereas in group II and III variants 
greater absorption intensity was observed for the FeIII-CN complex than the unligated FeIII state.  
Figure 4.21 shows the ligand binding curves for wtKatG, Q293A KatG (representing group I), 
IEEQ-A KatG and KatG?291 (representing group II) and all the other deletion variants 
(representing group III). Dissociations constants were determined by least-squares analysis of the 
absorption at 422 nm was fit to a single-ligand binding model. The KD values were for group I 
variants were similar to wtKatG but were significantly lower compared to the group II and III 
variants (Table 4.7). 
 
177 
  
 
310 350 390 430 470
0 .0 0 0
0 .0 1 5
0 .0 3 0
0 .0 4 5
0 .0 6 0
0 .0 7 5
0 .0 9 0
500 550 600 650 700
0 .0 0 0
0 .0 1 5
0 .0 3 0
0 .0 4 5
0 .0 6 0
0 .0 7 5
0 .0 9 0
408
424
A
540
642
W a v e l e n g t h  ( n m)
Ab
s
o
r
b
a
n
c
e
Ab
s
o
r
b
a
n
c
e
310 350 390 430 470
0 .0 0
0 .0 2
0 .0 4
0 .0 6
0 .0 8
0 .1 0
500 550 600 650 700
0 .0 0
0 .0 2
0 .0 4
0 .0 6
0 .0 8
0 .1 0
408 424
540
B
642
W a v e l e ng t h ( nm )
A
bs
o
r
ba
nc
e
A
bs
o
r
ba
nc
e
330 365 400 435 470
0 . 0 0
0 . 0 2
0 . 0 4
0 . 0 6
0 . 0 8
0 . 1 0
500 550 600 650 700
0 . 0 0
0 . 0 2
0 . 0 4
0 . 0 6
0 . 0 8
0 . 1 0
408
424
540 C
642
380
W a v e l e ng t h ( nm)
A
bs
o
r
ba
nc
e
A
bs
o
r
ba
nc
e
330 365 400 435 470
0 .0 0
0 .0 2
0 .0 4
0 .0 6
0 .0 8
0 .1 0
500 550 600 650 700
0 .0 0
0 .0 2
0 .0 4
0 .0 6
0 .0 8
0 .1 0
408
424
540
D
642
380
W a v e l e n g t h  ( n m )
A
b
s
o
r
b
a
n
c
e
A
b
s
o
r
b
a
n
c
e
 
Figure 4.20: Titration of wtKatG (A), Q293A KatG (B), KatG?291 (C), and KatG?288-293 (D) with potassium cyanide. Spectra 
were recorded 3 min after each addition of cyanide. All the reactions were carried out at 25 C in 100 mM phosphate buffer, pH 7.0 
 
178 
  
 
0 50 100 150 200
0 . 0 0 0
0 . 0 1 5
0 . 0 3 0
0 . 0 4 5
w t K a t G
Q2 9 3 A  K a t G
K a t G
? 291
K a t G
? LL2
K a t G
? 2 8 8 - 2 9 3
K a t G
? 2 9 1 - 2 9 2
I E E Q- A  K a t G
K a t G
? 2 9 0 - 2 9 3
[ K C N]  ( ? M)
?
A
222
 
Figure 4.21: Titration of wtKatG and group I, II, and III variants with potassium cyanide. 
The data point on the graph show ?A422 values as a function of the concentration of cyanide. The 
data were fit to a one binding site model to obtain KD values. All the reactions were carried out at 
25 C, in 100 mM phosphate buffer, pH 7.0 
 
 
 
 
 
 
179 
  
 
 
 
Table 4.7: Dissociation constants for cyanide binding of wtKatG and the variants obtained 
from equilibrium titration studies 
 
 KD (?M) 
WT KatG 1.4 ? 0.1 
Q293A KatG 1.5 ? 0.2 
KatG?291  8 ? 5 
IEEQ-A KatG 18.9 ? 0.6 
KatG?LL2 18 ? 2 
KatG?288-293 22 ? 1 
KatG?290-293 43 ? 2 
KatG?291-292 17.7 ? 0.7 
 
 
 
180 
  
4.3.4.2 Cyanide binding kinetics 
   Diode array spectra recorded upon reaction with KCN showed the similar changes as 
observed in cyanide titration experiments. The Soret absorption maximum was shifted from 408 
nm to 424 nm (isosbestic point 413) and a prominent new peak appeared at 540 nm in wtKatG 
and all three groups of variants. All the variants, including wtKatG, showed biphasic cyanide 
binding kinetics. In wtKatG and Q293A KatG (Figure 4.22 A), the biphasic cyanide binding 
included a rapid first phase whereas the first phase was much slower in group II (Figure 4.22 B) 
and III (Figure 4.22 C) variants.  
 The obtained pseudo first order rate constants for CN- binding by wtKatG and its variants 
increased linearly with KCN concentration (Figure 4.23). The apparent second-order rate 
constant (kon) for cyanide binding was calculated from the slope. For wtKatG and Q293A KatG 
the kon values were 4.3 x 105 M-1s-1 and 4.7 x 105 M-1s-1, respectively. Indeed, similar results 
were observed for all group I variants. In contrast, all the deletion variants as well as the 
quadruple substitution variant showed a three order of magnitude reduction in kon . The finite 
intercept gave the koff values, which are reported in Table 4.8. The koff / kon gave values for the 
dissociation constants (KD) of the cyanide complexes of each protein. KD values of 25 ?M were 
calculated for wtKatG and Q293A KatG whereas a much higher KD values (in mM) were 
obtained for group II and III variants (Table 4.8). 
 
 
 
181 
  
0 2 4 6 8 10
0 . 1 2
0 . 1 4
0 . 1 6
0 . 1 8
0 . 2 0
0 . 2 2
30
50
100
150
250
350
500
1500
A
T im e  ( s )
Abs
400
0 10 20 30 40 50
0 . 1 2
0 . 1 4
0 . 1 6
0 . 1 8
0 . 2 0
15
30
50
100
150
250
350
500
1500
B
T i m e  ( s )
A
bs
400
0 10 20 30 40 50
0 .1 2
0 .1 4
0 .1 6
0 .1 8
0 .2 0
15
30
50
100
150
250
350
500
1500
C
T im e  ( s )
Ab
s
400
 
Figure 4.22: Cyanide binding studies of Q293A KatG (A), IEEQ-A KatG (B), and KatG?288-
293. Panels A, B and C show the time course of formation of the ferri-cyano complex in the 
presence of varying concentrations (?M) of potassium cyanide at 400 nm. All reactions were 
carried out in single mixing mode at 25 C using 50mM phosphate buffer, pH 7.0  
 
182 
  
 
 
 
0 . 0 0 0 . 0 5 0 . 1 0 0 . 1 5 0 . 2 0 0 . 2 5 0 . 3 0 0 . 3 5
0
50
100
150
200
w t K a t G
Q2 9 3 A  K a t G
K a t G
? 2 8 8 - 2 9 3
I E E Q- A  K a t G
K a t G
? LL2
K a t G
? 2 9 1 - 2 9 2
K a t G
? 2 9 0 - 2 9 3
[ K C N]  ( m M )
k
o
b
s
(s
-1
)
 
 
Figure 4.23: Pseudo-first-order-rate constants for the formation of cyanide complex of 
wtKatG and variants as a function of cyanide concentration. Absorbance increase at 428 nm 
was fit to double exponential equation, and kobs values for the fast phase were plotted versus 
potassium cyanide concentration. All reactions were carried out in single mixing mode at 25 C 
using 50mM phosphate buffer, pH 7.0. 
 
 
 
 
183 
  
 
Table 4.8: kon, koff, and KD for cyanide binding by wtKatG, and its LL2 variants 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
KatG kon 
( 103 M-1s-1) 
koff 
(s-1) 
KD (?M) 
            WT KatG 434 ? 7 11 ? 1 25 
 
 
Q293A KatG 471 ? 11 12 ? 2 25 
 
 
IEEQ-A KatG 0.97 ? 0.07 0.27 ? 0.01 278 
 
 
KatG?LL2 0.018 ? 0.002 0.0189 ? 0.0003 952 
 
 
KatG?288-293 0.36 ? 0.03 0.254 ? 0.006 705 
 
 
KatG?290-293 0.6 ? 0.4  0.65 ?0.09 1083 
 
 
KatG? 291-292 0.7 ? 0.4 0.74 ? 0.09 1057 
 
184 
  
4.4. Discussion 
   KatG has a buried active site and more defined water matrix on the distal side of the 
heme compared to monofunctional peroxidases (102, 170-172). LL1 and LL2 help define a deep 
and narrow access channel to this buried active site. This architecture is typical for catalatic 
activity and its manipulation usually decreases the catalase but increases the peroxidase activity 
(43, 102). In this report, we introduced substitutions and deletions to the residues in the LL2 
structure of E.coli catalase-peroxidase to determine their influence on the active site architecture 
and function. Our previous studies on a variant obtained by deletion of the entire LL2 structure 
have shown that absence of LL2 results in a near complete loss of catalase activity with little 
effect on peroxidase activity (87). This suggests that LL2 in provides an architectural framework 
that is necessary for maintaining catalase-peroxidase bifunctionality.  
   Sequence alignments of different KatGs indicate a striking conservation of length in LL2 
rather than the identity of particular residues. Moreover, when the amino acid identity is 
conserved, with one exception (i.e., W297), glycine and proline are the conserved residues. 
When the amino acid character is conserved, it is in hydrophobicity. This is in contrast to the 
other large loop (LL1) where there is great variation in length, but at the same time, positions of 
sequence which are virtually invariant. Crystal structures indicate a point of contact between 
apex of LL2 and hydrophobic pocket of intersubunit C-terminal domain. The strict conservation 
of length and conformationally important residues suggests that the length is necessary to 
maintain this specific interaction. In this report, we explored the interaction of LL2 with the 
intersubunit C-terminal domain more closely to examine the role of this interaction on active site 
 
185 
  
function of KatG. We hypothesized that this interaction contributes to active site fine tuning for 
catalase activity and anticipated that any perturbation of this interaction would cause changes in 
heme coordination and consequently, activity of the protein.   
    Out of the six residues present at the apex of LL2, side chains Ile290 and Gln293 
protrude directly into a cavity provided by the C-terminal domain (Figure 4.1), whereas Glu291 
and Glu292 are both anticipated to point away from the pocket. Single and double substitutions 
of these amino acids produced variants that were catalytically and spectroscopicly similar to the 
wild-type enzyme. The catalase and peroxidase activities were more or less equivalent to the 
wild-type with apparent second-order rate constants around 106 M-1s-1 for catalase and 105 M-1s-1 
for peroxidase activities. Preservation of both the activities correlated well with unchanged 
spectroscopic characteristics. All of these variants contain predominantly hexacoordinate high-
spin heme as suggested by UV-Vis spectroscopy, and confirmed by MCD and EPR. Unchanged 
spectral features and activity profile might suggest that identity of the residue at the point of 
contact is not a particularly important criterion for maintaining the bifunctionality of KatG.  
   Conversely, deletion variants including the apex and limb residues differ drastically in 
catalytic and spectroscopic characteristics in comparison to the wild-type enzyme. Four apex 
deletion variants (KatG?288-293, KatG?290-293, KatG?291-292, and KatG?291) and two limb deletion 
variants (KatG?285/?294, and KatG?280/?299) were subjected to spectroscopic and kinetic analyses. 
The largest apex deletion variant i.e., KatG?288-293 had negligible catalase activity and with this 
came a notable shift in heme coordination environment toward the pentacoordinate state. These 
properties were essentially identical to the variant lacking the entire LL2 structure. Interestingly, 
 
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a similar pattern (i.e., shifts in heme coordination to pentacoordinate heme state and negligible 
catalase activity) was also observed for KatG?290-293 and KatG?291-292. The smallest apex deletion 
variant (KatG?291) and both the limb deletion variants retained around 20% of catalase activity 
but in terms of the heme coordination environment these resembled the rest of the deletion 
variants. All the deletion variants demonstrated decreased peroxidase activity with respect to 
H2O2 with very little effect on the ABTS-dependent peroxidase activity.   
   In contrast, the substitution variants had little effect, if any, on the catalase activity and 
consistent with these data, the spectroscopic features of these variants were comparable to those 
of the wild-type enzyme. It was not until, four of the apex residues were substituted with alanine 
(i.e., IEEQ-A KatG) that the effects similar to those seen for the deletion variants were observed. 
This quadruple substitution variant was dominated by pentacoordinate heme environment and 
retained only 6% of the catalase activity compared to wild-type enzyme. These observations 
emphasize on the fact that length conservation of LL2 is an important criterion to maintain the 
intersubunit interaction with C-terminal domain. This interaction, despite being 35? away from 
the active site, is essential for maintaining the bifunctionality.  
   In the absence of appreciable catalase activity, it is difficult to assess the specific reasons 
for the loss of catalytic function. Here, evaluation of remaining peroxidase activity and 
evaluation of small ligand (CN-) binding provide useful probes of important catalytic steps. 
Peroxidase kinetic parameters with respect to H2O2 provide insight into the ability to form 
compound I and other highly oxidized states necessary for catalysis. Kinetic parameters with 
 
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respect to the electron donor (e.g., ABTS) are informative to investigate electron transfer into the 
active site heme system.  
   The effect on H2O2-dependent peroxidase parameters was dramatic for all the deletion 
variants. The apparent second order rate constants with respect to H2O2 were two orders of 
magnitude lower than wild type enzyme. However, substantial rates of turnover (even in excess 
of wtKatG) could be observed at sufficiently high H2O2 concentration. This suggests a deficiency 
in the correct association of H2O2 in the active site leading toward its reduction to H2O. On the 
other hand, the kinetic parameters with respect to ABTS suggest that electron transfer to the 
heme by exogenous donors is unaffected and in some cases enhanced. 
   Attempts to use the stopped-flow to directly measure the formation of compound I were 
unsuccessful. The variants were insufficiently reactive at low peroxide concentrations, and at 
higher concentrations of peroxide, the variants showed signs of heme bleaching. However, small 
ligand binding to the ferric state, especially that of CN- mimics in many ways H2O2-dependent 
formation of compound I. We observed dramatically diminished rates of CN- binding for all the 
variants with LL2 deletions, both kon and koff decreased substantially with the net effect on KD 
being considerably larger for the deletion variants. Equilibrium titration experiments confirmed 
the same decrease in affinity for CN- as a ligand. Conversely, substitution variants (i.e., group I 
variants) showed CN- binding kinetics indistinguishable from wild type KatG. Equilibrium 
titration of CN- to determine KD for CN- for these variants confirmed the results of our stopped-
flow study. 
 
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   Interestingly, the diminished rates of CN- binding correlate with an increase in the 
proportion of heme in its pentacoordinate high-spin state and decrease in the hexacoordinate 
high-spin species. It is at first glance counterintuitive that the absence of a sixth ligand would 
diminish binding of CN- as a sixth ligand. After all, the presence of a pre-existing sixth ligand 
would be expected to compete with CN- for its binding site. However, it has been observed 
elsewhere that diminished rates of CN- binding by M. tuberculosis KatG go with increased 
proportions of the pentacoordinate state (70). Specifically, increased rates (1000-fold higher) of 
cyanide binding were observed in aged enzyme preparations due to higher proportions of 
hexacoordinate heme where a distal water molecule occupied the sixth position. On the contrary, 
freshly prepared enzyme dominated by pentacoordinate heme had lower affinity for cyanide as a 
heme ligand (70). Also, low temperature EPR spectroscopy and resonance Raman studies 
demonstrated favor ligand binding (cyanide or peroxide) to hexacoordinate heme. These results 
were suggested to provide evidence for the participation of a distal water molecule weakly bound 
at the sixth position of heme iron in the first phase of the reaction mechanism of KatG is 
important for the reaction of cyanide binding (70). 
   The distal water molecules have also been implicated in proton transfer within the distal 
cavity during the H2O2 - dependent compound I formation which is an initial step in both catalase 
and peroxidase turnover. QM/MM calculations addressing the compound I formation in HRP 
suggests an energetic advantage of specific distal water molecule in the active site (173). In 
Burkholderia pseudomallei KatG, mutation of Asp141, which is present at the acidic entrance 
channel and forms a hydrogen bond with Ile237, increases the KM for H2O2 oxidation. This was 
suggested to occur due to release of water molecule, which alters the solvent matrix in the 
 
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substrate channel for efficient H2O2 oxidation (143). So, it is reasonable to speculate that the 
displacement of distal water molecule from the sixth position in the deletion variants results in a 
loss of catalase activity by influencing H2O2 oxidation.  
   The first step of peroxidase turnover involves diffusion of H2O2 in to the active site and 
its deprotonation by the distal histidine acting as general base. Thus, the increased KM for H2O2 
in most of the deletion variants and two-order of magnitude lower apparent kcat/KM of all the 
deletion variants may suggests a deficiency in the correct association of H2O2 in the active site 
leading toward its reduction to H2O. This also suggests that deletion of parts of LL2 can disrupt 
the ordered matrix of oriented water dipole present in the narrow KatG access channel and this in 
turn can hamper the diffusion of H2O2 into the active site. These observations were further 
supported by cyanide binding experiments. Lower kon rates for cyanide binding for deletion 
variants also suggested the importance of distal water molecule in ligand access and binding. 
Moreover, lower KD values of wild-type enzyme and site-specific variants further supported that 
the hexacoordinate heme is the preferential species for cyanide binding.    
   In conclusion, the presented data suggest that length conservation of LL2 structure in 
KatGs is an important criterion with respect to maintaining the bifunctionality. Furthermore, the 
length conservation is highly essential to preserve the intersubunit interaction with the C-
terminal domain and this interaction despite of its distant location efficiently fine tune the active 
site structure of KatG to catalyze catalase and peroxidase reactions. Catalase activity parameters 
clearly demonstrate that deletion of any part of LL2 structure dramatically reduces / eliminates 
the catalase activity in contrast to site-specific substitutions. The lower apparent kcat/KM for 
 
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H2O2-dependent peroxidase activity along with diminished rates of catalase activity suggests that 
there are some difficulties in allowing the peroxide substrate access to the active site. This 
observation was further supported by cyanide binding studies which indicated that compound I 
formation is hampered in deletion variants. This may be due to disruption of the ordered network 
of water molecules especially at the distal part of the active site. The dominance of 
pentacoordinate high-spin heme species in all the deletion variants further helped us in 
suggesting that distal water molecule is displaced from the sixth ligand position which is 
preventing the efficient binding of peroxide substrate to the active site heme. Moreover, this 
work demonstrated that even the smallest deletion of LL2 structure was able to replicate the 
results as was seen in case of the enzyme lacking the entire LL2 structure. Thus, this work not 
only supported our previous findings that LL2 plays a major role in modulating the active site 
architecture and catalytic versatility of KatG but also provided sufficient evidences to support 
nature?s strategy to choose length conservation in this structure over the residue conservation. 
 
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Chapter Five: Summary 
 
Enzymes are nature's catalysts, tremendously accelerating the rates of a wide range of 
biochemical reactions, often with exquisite specificity. Clearly, nature is an accomplished 
enzyme engineer, and a central strategy employed by biological systems is the co-opting of 
existing structures for new catalytic purposes. One could envision several approaches for doing 
just that. For example, the direct substitution of active site amino acid residues, changing the 
geometric and electronic properties of the active site, could conceivably produce catalysts for as 
yet unaddressed reactions. Interestingly, since the advent of site-directed mutagenesis, 
biochemists the world over have tried just that approach. The result has been a near uniform 
failure. The virtually invariant result has been the loss of the existing catalytic abilities with no 
new added catalytic function. As such, it has been extraordinarily useful for understanding the 
mechanisms of existing enzymes, but uniformly poor for engineering new ones. 
This raises an important question: Is this the way nature has done enzyme engineering? 
While a definitive answer for all enzymes is not currently available, it is safe to say that in the 
case of catalase-peroxidases, the answer is clearly ?no?.  ith more than ample time to make 
substantial substitution of active site functional groups very little has changed between KatG and 
its fellow class I non-animal peroxidases (e.g., cytochrome c peroxidases), indeed, even 
comparisons to much more distantly related class II and III enzymes show very similar 
 
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active-site structures. Yet, KatG alone has catalase activity rivaling that of structurally unrelated 
monofunctional catalases. All the rest of the non-animal peroxidases have catalase activity at 
least three orders of magnitude lower. That ample time for active site substitutions has elapsed is 
easily evident from KatG?s C-terminal domain. Itself derived from a monofunctional peroxidase 
through gene duplication and fusion, the C-terminal domain shows substantial active site 
substitution such that it no longer even binds heme.    
This point to a strategy employed for the development of KatG as a catalase that can be 
understood in two ways. First, it represents the improvisation of a new mechanism for catalase 
activity. This improvised mechanism does not require substantial active site changes to 
accommodate it; rather this mechanism capitalizes on the strengths afforded by the active site 
structure as it is. In other words, the physical features of the peroxidases structure favor certain 
intermediates, interactions with substrates and chemical events. These are being used to 
accomplish catalase turnover in a new way. Second, a nearly identical active site indicates that 
where structural modifications have been made, these have primarily been carried out peripheral 
or external to the active. These must account for the subtle perturbation and fine adjustments 
necessary for the emergence of new catalytic function. 
The focus of the research described in this dissertation has been on the latter. In this 
regard, there are three structural components: a gene duplicated C-terminal domain, and two 
large loops- LL1 and LL2. The work in the Goodwin laboratory, including that described in this 
dissertation, demonstrates that all three structures play an important role in maintaining the 
bifunctionality of KatG enzymes. Moreover, this study demonstrates that KatG?s amenability to 
 
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structural fine-tuning makes it an ideal system for enzyme engineering. Based on the evidences 
provided in this dissertation and from the previous work in Goodwin lab, it is reasonable to 
suggest that catalase-peroxidases are fruitful models to study how enzyme functions are 
regulated by the fourth (i.e., global) structural level. Disruption of structural features unique to 
LL1 and LL2  (e.g., The W-Y-M covalent adduct or intersubunit noncovalent interactions, 
respectively) confirms that the changes to the structures distant to the active site would affect 
heme coordination state and catalytic properties of the enzyme.  
5.1. Effect on KatG catalytic versatility: Residue conservation of Large Loop 1 
The most obvious feature of the LL1 structure is its contribution of an invariant tyrosine 
to the W-Y-M covalent adduct. This seems rather a very small role for a structure which is 35-76 
amino acids long. Indeed, the LL1 structure is far more than is necessary to position tyrosine for 
participation in the adduct. KatG crystal structures show that this loop forms the part of substrate 
access channel and regulates the access of reducing substrates to the heme edge. Because of its 
location and its important role in supporting tyrosine this loop has been thoroughly investigated. 
Studies have shown that manipulation of the conserved residues of this loop present at the 
substrate entrance channel either diminishes or removes the catalatic activity of KatG but has 
varied effects on the peroxidase activity, ranging from no effect, decrease, or enhancement of the 
peroxidase activity. Also, the Goodwin lab has previously shown that removal of entire LL1 
causes complete loss of catalase activity and changes the rate limiting step to compound II 
reduction but increases the peroxidase activity compared to wild-type enzyme.  
 
194 
  
This suggested that there are some additional roles filled by this loop, which help to 
maintain the overall catalytic performance of KatG. The work in this dissertation has provided 
evidence in support of this hypothesis. The study involving the partial deletion of LL1 (KatG?209-
228) and its comparison to Y226F KatG and KatG?LL1, helped elucidate not only the mechanism 
for loss of catalase activity but also the substantial enhancement of peroxidase activity. It has 
become clear that the LL1 structure is essential for maintaining the structural integrity of KatG, 
as deletion of the entire structure makes the protein insoluble during expression. Furthermore, 
this loop favors catalase turnover by obstructing direct access of reducing substrates to the heme 
center. Simultaneously, it rescues the enzyme from inactivation which would normally arise due 
to reducing substrate unavailability, by cleverly providing a tyrosine for the formation of the W-
Y-M adduct. This has been proposed to facilitate the release of molecular oxygen to complete a 
novel catalatic cycle.  
The effect of the LL1 structure on the catalytic activity of KatG would be interpreted as 
uncovering a mutual competition between catalase and peroxidase turnover. However, recent 
work from the Goodwin laboratory, involving mechanistic studies of KatG catalytic cycle in M. 
tuberculosis have suggested that peroxidase activity acts as a synergistic mechanism for catalase 
activity. This mechanistic study forces one to look at the peroxidase and catalase activities of 
KatG from a whole new perspective. Up to now, the general consensus of the fields is that both 
the activities are competitive in nature and this has led to the classical representation of catalase-
peroxidase mechanistic scheme of KatGs, which is just an overlay of individual catalase and 
peroxidase cycles. And more so, the conditions used to measure one activity (e.g., pH, buffer 
conditions, availability of substrates, etc.) eliminate contribution from the second. This suggests 
 
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that at any given point of time enzyme is favoring one activity over the other. Thus, in the 
absence of one activity (e.g., catalase) no change or even enhancement of the other (e.g., 
peroxidase activity) is seen which again emphasizes the competitiveness between the two 
activities. However, the data from recent mechanistic studies suggested that the presence of 
peroxidase activity or more importantly the presence of reducing substrates enhance the catalase 
activity instead of diminishing or eliminating it. This has led to the new hypothesis that the 
peroxidase activity has been folded into making catalase activity of KatG operate by a novel 
mechanism producing more efficient catalyst over a broader range of conditions. The work, 
though in its initial stages, has provided enough data to reveal the devotion of peroxidase 
machinery towards enhancement of the catalase activity, especially under conditions where 
catalase activity is assumed to be absent. Under this paradigm, the presence of LL1 structure and 
its constraints for the electron donors to enhance the catalase activity can help us envision the 
presence of peroxidase activity as a catalase activity enhancer. 
5.2. Effect on KatG catalytic versatility: Length Conservation of Large Loop 2 
Previously it has been shown that LL2 is critical for the bifunctionality of KatG. Deletion 
of entire LL2 caused a 99% loss of catalase activity along with dominance of pentacoordinate 
high spin heme species over the hexacoordinate that dominates in the wild type enzyme. This 
showed that removal of entire loop has a profound impact on the KatG catalytic activity and its 
absences make this bifunctional enzyme like monofunctional peroxidases.  
 
196 
  
To explore how LL2 affects KatG function, the effects of the disruption of this peripheral 
structure were examined. Sequence comparison of 20 widely phylogenetically dispersed KatGs 
showed a remarkable length conservation of LL2 but relatively little sequence conservation. The 
LL2 is ~35 amino-acid long, starts and ends near the active site, and spans between F and G 
helices. Most importantly, the apex of LL2 forms a unique intersubunit interaction with the C-
terminal domain. Deletion of the entire LL2 (KatG?LL2) changed the coordination state of the 
enzyme to pentacoordinate ferric heme and rendered the enzyme catalase negative. KatG?LL2 
showed mixed populations of high spin ferric heme with 77% rhombic contribution and 23% 
axial contribution in contrast to 99% axial contribution in wild type enzyme. This suggested that 
LL2 preserves the catalatic activity of KatG by maintaining the proper active site architecture. 
This role of LL2 was further strengthened by the fact that disruption of intersubunit interactions 
between LL2 and C-terminal domain were sufficient to duplicate the results of KatG?LL2 . indeed, 
just deleting two amino acid residues at the point of interaction. Even the elimination of one 
amino acid out of 35 reduced catalase activity by 80%. The importance of length conservation of 
this loop was strengthened by the fact that deletion variants had greater impact on overall 
catalytic activity and heme coordination of KatG compared to the site specific variants involving 
the apex residues. 
The deletion variants showed the heme  species population mixture of KatG?LL2. This 
was confirmed by UV-Vis, MCD, and EPR spectroscopy. In contrast, site-specific variants 
showed higher percentage of hexacoordinate high spin heme similar to the wild-type enzyme. 
The steady state kinetics showed that deletion variants retained little if any catalase activity 
whereas site-specific variants were more or less catalase active. The effect on peroxidase activity 
 
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of deletion variants was variable. The apparent kcat/KM with respect to H2O2 was diminished by 
two orders of magnitude with very little effect on ABTS- dependent parameters. The rates of 
cyanide binding were dramatically diminished in deletion variants whereas the rates were 
indistinguishable from wild-type enzyme for the substitution variants.  
In conclusion, the dominance of pentacoordinate high-spin heme species, near absence of 
catalase activity, decreased rates of H2O2-dependent peroxidase activity, and finally diminished 
rates of cyanide binding in all the deletion variants suggested that length conservation of LL2 is 
an important criterion to maintain the distant intersubunit interaction. And, this intersubunit 
interaction between LL2 structure and intersubunit C-terminal domain preserves the catalatic 
activity to KatG by retaining the proper active site architecture. 
5.3. Conclusions 
In summary, this research has helped to identify how nature has gone about engineering a 
new catalytic activity from an old structural scaffold. The importance of global structures in 
balancing the catalytic performance of KatG suggests that nature?s decision to manipulate 
KatG?s global structure has made this enzyme a better catalyst and opened up many doors for 
engineering a new catalyst. Nature?s strategy to use a peroxidase scaffold and manipulate 
through the action of peripheral structures to build a new function is an elegant one. On one 
hand, residue conservation in LL1 has provided an excellent gateway to enhance the catalase 
function by using peroxidase machinery. On the other hand, length conservation in LL2 has 
maintained a necessary active-site structural framework by interacting with the  C-terminal 
 
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domain of another subunit some 35? away. This is the kind of interaction that would typically be 
overlooked due to its distant location. But this research has clearly shown that this distant 
interaction is crucial in order to maintain the bifunctionality of KatG. This research points 
toward the necessity of understanding peripheral structures and distant protein-protein 
interactions for controlling catalytic function.  
 
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