Site-Directed Mutagenesis and Ruthenium Labeling of Oxalate Decarboxylase

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Site-Directed Mutagenesis and Ruthenium Labeling of Oxalate Decarboxylase
DeMason, Timothy S.
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Oxalate is a toxic dicarboxylic acid that is decomposed into carbon dioxide and formate by Bacillus subtilis Oxalate Decarboxylase (OxDC). Each monomer of OxDC contains two manganese ions, one at the N-terminal end and one at the C-terminal end. Evidence suggests that the C-terminal manganese plays a functional role in the mechanism of catalysis, which can be further investigated by studying a ruthenium labeled OxDC. A K375C/C383A OxDC mutant was generated, which places a cysteine close to the C-terminal manganese for labeling with the thiol reactive ruthenium compound [Ru(bpy)2(IA-phen)]2+. Marcus theory calculations predict a tunneling time of 660 ns between the ruthenium and C-terminal manganese ions. The K375C/C383A OxDC mutant displayed typical wild type Michaelis-Menten kinetics, with KM = 10 ± 2 mM and kcat = 90 ± 13 s-1. Attempts at labeling K375C/C383A OxDC with [Ru(bpy)2(IA-phen)]2+ were unsuccessful. The failure of the labeling reaction appears to be due to an inability of [Ru(bpy)2(IA-phen)]2+ to react with K375C/C383A OxDC rather than nonspecificity of the reaction. Future labeling with other thiol reactive ligands should be attempted. ( en )
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Awarded Bachelor of Science, summa cum laude, on May 8, 2018. Major: Chemistry. Emphasis/Concentration: Biochemistry
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College or School: College of Liberal Arts and Sciences
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Advisor: Alexander Angerhofer. Advisor Department or School: Department of Chemistry

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Site Directed Mutagenesis and Ruthenium Labeling of Oxalate Decarboxylase Timothy DeMason Undergraduate Honors Thesis Spring 2018 Department of Chemistry, University of Florida


2 Contents List of Abbreviations ................................ ................................ ................................ .................... 3 Abstract ................................ ................................ ................................ ................................ ......... 4 Introduction ................................ ................................ ................................ ................................ ... 5 Methods ................................ ................................ ................................ ................................ ......... 1 1 Results and Discussion ................................ ................................ ................................ ................. 1 6 Calculation of Ru Mn Electron Transfer ................................ ................................ ....... 1 6 Mutagenesis of K375C/C383A ................................ ................................ ......................... 1 8 S ynthesis of Ru OxDC ................................ ................................ ................................ ..... 2 2 Conclusion ................................ ................................ ................................ ................................ .... 2 5 Acknowledgements ................................ ................................ ................................ ....................... 2 6 References ................................ ................................ ................................ ................................ ..... 2 6


3 List of Abbreviations bis tris 2 b is(2 hydroxyethyl)amin o 2 (hydroxymethyl) 1,3 propanediol bpy bipyridine DMSO dimethyl sulfoxide DTT dithiothreitol E. coli Escherichia coli FDH F ormate D ehydrogenase IA phen 5 iodoacetamido 1,10 phenanthroline IPTG i D 1 thiogalactopyranoside LRET Long Range Ele ctron Transfer mM mili Molar micro Molar MS Mass Spectrometry NAD + n icotinamide a denine d inucleotide OxDC Oxalate Decarboxylase PCET Proton Coupled Electron Transfer PCR Polymerase Chain Reaction PDB Protein Data Bank Ru OxDC Ruthenium Modified Oxalate Decarboxylase Tris 2 a mino 2 (hydroxymethyl) 1,3 propanediol WT Wild Type Oxalate Decarboxylase


4 Abstract Oxalate is a toxic d icarboxylic acid that is decomposed into carbon dioxide and formate by Bacillus subti lis Oxalate Decarboxylase (OxDC). Each mo nomer of OxDC contains two manganese ions, one at the N terminal end and one at the C terminal end Evidence suggest s that the C terminal manganese plays a functional rol e in the mechanism of catalysis, which can be further investigated by studying a rut henium labeled OxDC. A K375C /C383A OxDC mutant was generated which places a cysteine close to the C terminal manganese for labeling with the thiol reactive ruthenium compound [Ru(bpy) 2 (IA phen)] 2 + Marcus theory calculations predict a tunneling time of 6 60 n s between the ruthenium and C terminal manganese ions The K375C/C383A OxDC mutant displayed typical wild type Michaelis Menten kinetics, with K M = 10 2 mM and k cat = 90 13 s 1 Attempts at l abeling K375C/C383A OxDC with [Ru(bpy) 2 (IA phen)] 2 + wer e unsuccessful. The failure of the labeling reaction appears to be due to an inability of [Ru(bpy) 2 (IA phen)] 2 + to react with K375C/C383A OxDC rather than nonspecificity of the reaction. Future labeling with other thiol reactive ligands should be attempte d.


5 Introduction 1 Oxalic acid is produced in plants through several biochemical pathways including the activity of glyoxalate oxidase and isoci trate lysase. 2 3 Oxalate is known to precipitate in the presence of divalent cations. 1 Calcium oxalate is one of the more important salts since it is rel atively insoluble and is present in about 60% of kidney stones 4 Developing ways of reducing the degree of oxa late accumulation in the kidneys is of medical importance. The buildup of calcium oxalate deposits can also cause pro blems in the manufacturing of paper 5 6 Oxalate Decarboxylase (OxDC) is an oxalate degrading enzyme found in Bacillus subtilis and other organisms. The enzyme catalyzes the degr adation of oxalate in to formate and carbon dioxide in 99.8% of turnovers (decarboxylase pathway) and two equivalents of carbon dioxide and hydrogen peroxide in the rest (oxidase pathway) as shown in Scheme s 1 A and 1B 1 Scheme 1 : A) Degradation of oxalate into carbon dioxide and formate and B) of oxalate into carbon dioxide and hydrogen peroxide. Native OxDC exists as a homo hexamer formed from a dimer of trimers (see Figure 1 C). The struct ure of the monomer is shown in Figures 1 A and 1 barrel domains one at the N terminal end ( shown in green in Figures 1 A and 1 B) and another at the C


6 terminal end ( shown in blue in Figures 1 A and 1 B). Inside each barrel rests a manganese ion bound to three histidines and one glu tamate, leaving two sites free for other small ligand s Figure 1 : The crystal structure of Bacillus subtilis OxDC showing the manganese ion s in purple. A) Monomer structure as viewed from the side. B) Monomer viewed from the top. C) Hexamer viewed from the top with one trimer in blue and the other in orange. These figures were generated in PyMOL using the 1UW8 P D B file. The mechanism of OxDC catalysis of oxalate is still not completely understood. Figure 2 illustrates the current proposed mechanism for catalysis. First mono protonated oxalate and di oxygen bind to the N terminal manganese with the di oxygen generating Mn 3+ 1 Then glutamate 1 62 removes the remaining acidic proton from oxalate, while simultaneously an ele ctron is transferred from oxalate to the manganese i.e. proton coupled electron transfer (PCET) 1 Heterolytic cleavage of the oxala te carbon carbon bond follows, liberating carbon dioxide and producing a carbon dioxide radical anion still bound to the manganese 1 Finally, PCET occurs again, reducing the carbon dioxide radical anion while oxidizing the Mn 2+ to A ) B) C)


7 regenerate Mn 3+ Concurrently, g lutamate 162 protonat es the carbon of the carbon dioxide ra dical anion to form formate 1 A problem with this mechanism is that it places a dioxygen radical and carbon dioxide radical anion in close proximity These two radicals are expected to react to form peroxycarbon ate (HCO 4 ) wh ich may then decompose with proton uptake to produce hydrogen peroxide and carbon dioxide as is the case in the oxidase pathway. 7 However, the products of the oxidase pathway are only observed in 0.2% of turnovers. 1 Figure 2 : Current proposed mechanism for OxDC. The N termin al manganese binds oxalate (and oxygen as a co catalyst ) while it cycles between Mn 2+ and Mn 3+ From ( 9 ) with permission from Elsevier. While th e C terminal manganese was originally thought to only play a structural role, further investigation into the OxDC mechanism suggest s that the C terminal manganese may play a role in catalysis. The presence of a stacked tryptophan dimer between the N and C terminal manganese of adjacent monomers suggests that electron transfer between the two manganese ion s is possible. Substitution of tryptophan 96 and tryptophan 274 with


8 phenylalanine or tyrosine leads to significantly reduced catalytic capability indica ting their importance in catalysis 1 8 Additio nally, EPR spin trapping studies of a flexible lid mutant suggest that the carbon dioxide radical anion s and superoxide radicals are produced in separa te locations. 9 Since o xalate is known to bind at the N terminal manganese perhaps oxygen bind s to the C terminal manganese. 1 A new mechanism for OxDC catalysis is shown in Figure 3 proposing the C terminal manganese site as the temporary electron sink through the use of long range electron transfer (LRET) 10 10 Figure 3 : New proposed mechanism for OxDC, inc ludin g the possibility of long range electron transfer assuming oxalate binds in a bi dentate fashion From ( 10 ).


9 Selective oxidation of the C terminal manganese could be used to further study the possibility of LRET. Ruthenium m odified proteins have been used previously to study the electron transfer properties of metalloproteins such as azurin, cytochrome c and myoglobin 11 These modified proteins employ the use of the so ique shown in Scheme 2 A A ruthenium(II) diimine complex such as tris (bipyridine)ruthenium(II), is excited of light an oxidant or reductant to prod uce ruthenium(III) o r ruthenium (I), respectively. 12 The oxidized or reduced ruthenium can then either remove or inject an electron from a suitable nearby target. 12 Scheme 2 : A ) General f lash quench scheme showing both the oxidation and reduction route s with reduction potentials B ) Oxidation scheme proposed for OxDC using a small quencher Q. Modified from ( 12 ) with permissi In the case of OxDC, bis(bipyridine)(5 iodoacetamido 1,10 phenanthroline)ruthenium(II) ( i.e [Ru(bpy) 2 (IA phen)] 2+ ) can be used to oxidize the C terminal manganese (see Scheme 2B). The iodoacetamide group of [Ru(bp y) 2 (IA phen)] 2+ is expected to react with reduced, surface accessible cysteines though an S N 2 mechanism as shown in Scheme 3 12 This reaction covalently links the ruthenium complex to the protein. The ruthenium modified OxDC ( Ru OxDC) can then be studied to further explore the role of the C terminal manganese in any electron transfer steps. A) B)


10 Scheme 3 : Reaction of [Ru(bpy) 2 (IA phen)] 2+ with a cysteine residue. In order to label OxDC with this ruth enium complex, a cysteine needs to be introduced at the C terminal site. Furthermore, all other cysteines need to be removed to promote selective labeling at the C terminal site. Previously a C383A OxDC mutant was created by Dr. Umar Twahir in the Angerho fer Lab This mutant serve d as the starting place for this project since the only cysteine in the OxDC sequence has been removed.


11 Methods Mutagenesis was p er formed using a Q5 Site Directed Mutagenesis Kit from New England Biolabs. Briefly, pr imers for two separate mutants were designed using NEBaseChanger to incorporate the cysteine TGC codon at the appropriate site (see Table 1 ) and purchased from I ntegrated DNA Technologies Table 1 : Site Directed Mutagenesis Primers Primer K37 5 C FWD TTCAAAAGAAtgcCACCCAGTAGTGAAAAAG K37 5 C REV AGCACATCAGTAAAGTCTTTG A341C FWD CGACCATTATtgcGATGTATCTTTAAACCAATG A341C REV TCTTTGAAGATTTCTAAAAAGAC Then, a mix of 0.8 ng /mL template DNA (pET 32a plasmid containing the C383A mutation) 0.5 of both forward and reverse primers, and Q5 Hot Start High Fidelity 1 X Master Mix, was prepared and placed in a thermocycler for 25 cycles. Before the cycling began, the samples were initially denatured by heating at 98 C for 30 s. The cycles consisted of a 10 s denaturation step at 98 C followed by a 30 s annealing step at either 5 6 58, or 61.5 C and finally an extension step at 72 C for 3 min A final extension step was performed again at the end of the 25 cycles followed by a holding period of 5 to 1 0 min at 4 C The PCR product was then incubated in a KLD mix (contains a kinase, ligase, DnpI) to phosphorylate digest the methylated template and ligate the PCR product. NEB 5 alpha c ompetent E. coli cells were then transformed with 30 s of heat shoc k at 42 C The transformed cells were st r eaked onto LB agar plates treated with 50 ng/ L ampicillin and grown overnight. A colony was then selected and g r own in an overnight culture for plasmid purification the following day. The miniprep was performed us ing a Wizard


12 Plus SV minipreps DNA Purification System. Some plasmid was sent to GENEWIZ ( 115 Corporate Blvd, South Plainfield, NJ 07080 ) for sequencing, and some was used to transform BL21 (DE3) c ompetent E. coli cells which were grown for a glycerol stoc k and stored at 80 C for future protein expression Protein expression and purification was performed using protocols established in the literature 9 First, a small amount of stock E. coli cells of the desired mutant was gr own in an overnight culture using ampicillin treated LB media ( 50 n g/ L ampicillin, 5 g/L yeast extract, 10 g/L tryptone, 85 mM NaCl) at 37 C Then, a sample of cells from the overnight culture w as grown in 3 L of ampicillin treated LB media at 37 C unti l OD 600 =0.5. At that point, the cells were heat shocked at 42 C for 10 minutes and 5 mM Mn Cl 2 and 0.8 mM i D 1 thiogalactopyranoside ( IPTG ) were introduced into the cultures to induce expression of OxDC After four hours of expression, the cells were pelleted and stored at 80 C. When purification was ready to be performed the cell pellets were thawed and resuspended in 40 mL of lysis buffer ( 2 10 mM imidazole pH 7.5 ) before being sonicated The lysed cells were pelleted down and the supernatant p oured into a purification column containing 5 mL of washed Ni NTA resin and shaken at 4 C for 2 hours. Then, the column was drained of the supernatant and 50 mL of wash buffer (50 mM T ris, 500 mM NaCl, 20 mM imidaz ole pH 8.5 ) was passed through the column at 4 C followed by 40 mL of elution buffer (50 mM pH, 500 mM NaCl 250 mM imidazole 8.5 Tris ) F ractions of every elu a t e were collected every 5 10 mL. These fractions were dialyzed overnight in 2 L of st orage bu ffer (50 mM Tris 500 mM NaCl pH 8.5 ) concentrated the next day following treatment with 50 mg/mL Chelex resin to remove free metals Aliquots of enzyme were finally flash frozen and stored at 80 C


13 Enzyme kinetics of C383A and K375C/C383A OxDC mutants w ere studied by using an end point formate dehydrogenase (FDH) coupled assay making use of the reaction shown in Scheme 4 to measure the amount of formate produced Scheme 4 : Reduction of NAD + by formate, catalyzed by FDH The assay was performed by mixing a 5 pH 4.2 poly buffer (piperazine, tris, bis tris, and acetate 50 mM each ) 500 mM NaCl, 0.5 mM ortho phenylenediamine 0. 004 % (m/v) triton X, and concentrations of oxalate varying bet ween 2 and 100 mM The samples were react ed 50 mM pH 7.8 phosphate buffer 1. 5 mM NAD + and 0. 0004 % (m/v) FDH and incubate d over night at 37 C. Finally, absorbance reading s were taken at 340 nm and the concentration of formate produced was calculated using a standard curve obtained on the same day. The labeling of the K375C/C383A mutant was performed in two slightly different cond itions B oth were similar to a previously described protocol. 12 First, a 2 mL sample of 2.2 mg/mL K375C/C383A OxDC was reduced at pH 8 with 5 mM dithiothreitol ( DTT ) for 30 min at 4 C Then, the DTT was dialyzed out of the sam ple at 4 C for 2 hours using 2 L of pH 8 storage buffer A solution of ruthenium label was prepared by dissolving b etween 1 and 2 mg of [Ru(bpy) 2 (IA phen)](PF 6 ) 2 (purchased from Santa Cruz Biotech) was in 1 mL DMSO and further diluted with 1 mL pH 8 stora ge buffer All 2 mL of the ruthenium label solution was subsequently added to the reduced OxDC to initiate the reaction shown in Scheme C The mix


14 was shaken for 4 hours at 4 C in the dark. Finally, the end product was dialyzed overnight to remove excess label and the sample was concentrated. The labeling process was also performed at 25 C A 1.5 mL sample of 4.0 mg/mL K375C/C383A OxDC was reduced with 5 mM DTT at pH 8 for 30 min at 25 C Then, the DTT was dialyzed and the ruthenium label solution prepa red as previously described. All 2 mL of the ruthenium label solution was subsequently added to the 1.5 mL of the reduced OxDC to initiate the labeling reaction. The mix was shaken for 4 hours at 25 C in the dark. Finally, the end product was washed with 20 mL of pH 8 storage buffer to remove the excess ruthenium label and subsequently concentrated. Trypsin digest and mass spectrometry analysis was performed under the direction of Dr. Kari Basso of the UF Department of Chemistry. Briefly, s amples of prot ein were processed via SDS PAGE on a 4 15% acrylamide gradient gel from Biorad The band corresponding to OxDC was cut from the gel washed with nanopure water, and dehydrated with 1:1 v/v acetonitrile and 50 mM (NH 4 )HCO 3 The gel band was then rehydrated with 12 ng/ml sequencing grade trypsin in 0.01% ProteaseMAX Surfactant and then overlaid with 40 L of 0.01% ProteaseMAX S urfactant and 50 mM (NH 4 )HCO 3 and gently mixed for 1 hour. The digestion was stopped with the addition of 0.5% trifluoroacetic acid Next, nano liquid chromatography tandem mass spectrometry (Nano LC/MS/MS) was performed on a Q Exactive HF Orbitrap mass spectrometer equipped with an EASY Spray nanospray source operated in positive ion mode. The LC system used was an RSLCn ano. The mobile phase A was 0.1% formic acid and acetic acid in water and the mobile phase B was aceto nitrile with 0.1% formic acid in water First, 5 L of the sample was injected onto a 2 cm C18 column and washed with mobile phase A. The injector port wa s switched to


15 inject and the peptides were eluted off the trap onto a 25 cm C18 column for chromatographic separation. P eptides were eluted directly off the column into the LTQ system using a gradient of 2 80% B with a flow rate of 300 nL/min. T he total ru n time was 60 minutes The EASY Spray source operated with a spray voltage of 1.5 k V and a capillary temperature of 200 o C. The scan sequence of the mass spectrometer


16 Results and Discussion Calculation of Ru M n Electron Transfer In order to predict whether the ruthenium complex will oxidize the C terminal manganese, the following Marcus equation (Equation 1) can be used to predict the rate constant of electron transfer, k ET : 13 (1) where is the reorganization parameter, H AB is the electr onic coupling between reactants and products and G is the reaction driving force 11 The electronic coupling factor, H AB can be approximated by using the regression shown in Figure 4 The upper limit of the Mn Ru distance can be estimated by adding the distance between the Mn and the sulfur of cysteine 375 and the radius of the [ Ru(bpy ) 3 ] 2+ c omplex The Mn S distance w as estimated to be 11.1 from using the crystal structure OxDC (P D B 1UW8) to find the distance between the Mn and the carbon of lysine 375 The diameter of [Ru(bpy) 3 ] 2+ was reported as 5.4 in the literature. 14 Figure 4 : Plot of H AB vs donor acceptor distance (R DA ) for thermal (magenta) and op tical (blue) intramolecular ET. From ( 11 ) with permission from the ACS.


17 Thus, the M n Ru distance was taken to be approximately 16.5 so by using Figure 4 H AB is approximately 0.1 cm 1 The value of was estimated to be 0 .8 e V in accordance the t ypical reorganization parameter for protein ET processes 11 The driving force of the reaction can be calculated using the reduction potential of [Ru(bpy) 3 ] 2+ and the oxidation potential of a OxDC manganese model complex as estimates. 15 16 Ru 3+ + e 2+ E = +1.3 V (vs NHE) Mn 2+ 3+ + e E = 0.73 V (vs NHE) Taking G = 0.57 eV along with the other previous stated parameters and using Equation 1, we find that k ET = 1 5 10 6 s 1 with a tunneling ET = 660 n s. Because ET is on the order of hundreds of nano seconds it is expected that the Mn Ru electron transfer will occur since electron transfer has been ob served in similar ruthenium modified proteins with ET on the order of milliseconds or faster 11 Both the values of the manganese reduction potential and the Mn Ru distance are rough estimates. Modest change s in either of these parameters could have noticeable impacts o n the rate of electron transfer. To minimize the tunneling time it may be n ecessary to find a photosensitizer that produces a driving force of G T he bipyridines can be modified by adding either electron donating/withdrawing groups to alter the reduction potential of the ruthenium complex. 17 Furthermore ruthenium can be substituted with another d 6 metal such as rhenium(I) or osmium(II) to further adjust the reduction potential. 18


18 Mutagenesis of K375C/C383A Based on the crystal structure of OxDC alanine 341 and lysine 375 appeared to be the closest surface accessible re s idue s to th e C terminal manganese with distances of 10.1 and 11.1 , respectively (see Figure 5 ). These two sites were chosen as candidates for mutation to a cysteine. Figure 5 : Location of alanine 341 and lysine 375 ( orange ) relative to the C terminal manganese wi th distances between the residue and the manganese given too. This figure was generated in PyMOL using the 1UW8 P DB file. Site directed mutagenesis was then preformed using the non overlap extension method as described in the Methods section A sample of the unligated PCR product was analyzed using agarose gel electrophoresis stained with ethidium bromide which is displayed in Figure 6 As see n in the gel, bands for the K375C mutation are clearly visible but the bands for the A341C mutation are not. This indicated that the PCR was not successful for the A341 mutation but was successful for the K375C mutation.


19 Figure 6 : Agarose gel of PCR product. Lanes 1 3 contained samples from the A341C mutation at 56, 58, and 61.5 C, lanes 4 6 contained samples from the K375C mutation at 56, 58, and 61.5 C and lane 9 contained a ladder Sa mples of the K375C plasmid were sequenced by GENEWIZ using the Sanger sequencing method The sequence of the forward strand did not conclusively show the mutation, but the sequen ce of the reverse strand did. Figure 7 shows the reverse complement of a section of the reverse strand sequencing data. T his region displays the same sequence as the forward primer indicating that the mutation was successfully incorporated into the synthe sized DNA. Figure 7 : Select DNA sequencing results showing the sequence of the reverse complement to the reverse strand. Figure generated by GENEWIZ software using the trace file from GENEWIZ for the reverse sequence. 1 2 3 4 5 6 7 8 9 10


20 Additionally, the sequencing data w as compared to the sequence of OxDC (with the cysteine mutation included) using the NCBI Nucleotide BLAST. B oth the forward and reverse sequence matched the comparison sequence for the bases sequenced The kinetics of the C383A and K375C/C383A mutants w er e studied using the FDH coupled assay described earlier. Lineweaver Burk plots (normalized of Mn content) were constructed for each mutant and are shown in Figures 8 and 9 Figure 8 : Lineweaver Burk plot constructed for the C383A mutant. A linear regress ion was performed generating a line given by y= 0.14266 9x+ 0.0099 9 with R 2 =0.959 Figure 9 : Lineweaver Burk plot constructed for the K375C/C383A mutant A linear regression was performed generating a line given by y= 0.07982 5x+ 0.00802 with R 2 =0.983. 0 0.02 0.04 0.06 0.08 0.1 0.12 0.14 0.16 0 0.1 0.2 0.3 0.4 0.5 0.6 v 1 1 ) [oxalate] 1 (mM 1 ) 0 0.01 0.02 0.03 0.04 0.05 0.06 0.07 0 0.1 0.2 0.3 0.4 0.5 0.6 v 1 1 ) [oxalate] 1 (mM 1 )


21 Fro m these Lineweaver Burk plots, values of K M and k cat were calculated (normalized for Mn content) and are displayed in Table 2 along with a range of values for wild type (WT) OxDC reported shown for comparison Table 2 : Michaelis Menten Kinetics of Produced Mutants When accounting for the margin of error, the v alues for K M and k cat fall within the range of values reported for WT OxDC in the literature This indicated that neither mutation has a significant effect on the kinetics of the enzyme. Thus, the K375C/C383A mutant appears to be a suitable candidate for labeling with [Ru(bpy) 2 ( I A phen)] 2+ K M (mM) k cat (s 1 ) C383A 14 4 73 15 K375C/C383A 10 2 90 13 WT 7 5 1 28 1 WT 19 6.6 0.6 71 6 WT 20 12 3 158 13


22 Synthesis of Ru OxDC The labeling of K375C/C383A was performed as described in the Methods section. Initially, the reaction was performed at 4 C. A sample of the end product was then digested with trypsin and analyzed by mass spectrometry (MS) as describe d previously to identify if either the full ruthenium label or the p henanthroline ligand was bound to cysteine 375 The MS results did no t show a peak corresponding to a fragment with either label Additionally, a sample of the end product was washed with 15 mL of pH 8 storage buffer. The prominent yellow color produced by the ruthenium complex gradually disappeared until the sample became colorless, further indicating that the label wa s not bound to the enzyme. It was hypothesized that the cysteine may not be surface accessible at 4 C so an t of free cysteines at 4 C and at 25 C. A negligible amount of free cysteines was found at 4 C, but at 25 C there were 0.6 free cysteines per monomer. The refore, the labeling process was reperformed at 25 C. The M S results of the unpurified product of this reaction also did not show a peak corresponding to the labeled cysteine 375 fragment. The end product of this reaction was also washed as described in the Methods section to determine if the yellow orange color would gradually fade as was the case for the 4 C sample. After washing with 20 mL of pH 8 storage buffer, the sample was a lighter ye llow orange color and the eluate was colorless, suggesting that some ruthenium could be bound to the enzyme. Interestingly, during the washing process some unknown orange precipitant was formed. To further investigate why the reaction did not occur as exp ected, a potential structure of the Ru OxDC protein was created in Py MOL using the mutagenesis and bond fusing features to perform the K375C mutation and attach [Ru(bpy) 2 ( I A phen)] 2+ to the sulfur (Figure 10 A)


23 Figure 10 : A) Estimated location of the [Ru(bpy)(IA phen)] 2+ complex relative to the C terminal Mn B) R esidues within 6 displayed in orange Figures generated in PyMOL using the 1UW8 PDB file After the structure was created, the residues within 6 were displayed to check for any steric interference. Figure 10 B shows this display and reveals that there is sufficient space for the ruthenium compound to sit above the opening of the C barrel. It is possible that the labeling reaction did not occur as expected because the ruthenium label was binding to another site on the enzyme. One possibility is that the label was reacting with a lysine residue. In order to determin e if this was the case the MS data was searched to find a fragment corresponding to labeled lysine 20 which was found to be surface accessible and the most nucleophilic in crosslinking expe r iments. 8 Again, the MS results did not show a labeled fragment. Another possibility is that the unprotontated nitrogen of a h istidine residue could react s iodoactamid e group H owever this is not likely since histadine 376 is surface A) B )


24 accessible and is adjacent to cysteine 3 75 but MS data showed that fragment was not labeled. While it does not appear that histidine reacts with the iodoacetamide group, it could potentially coordinate with the ruthenium replacing one of the other ligands This would likely occur at the C termi nal histidine tag where there are 6 histidines. If this were the case then there should be some free IA phen in the solution which should then bind to the protein but this is not observed in the MS data. Perhaps it would be best to use a different function al group to attach the ruthenium compound to OxDC. While there have been methods developed to attach similar ruthenium compounds to a lysine or to coordinate them with a histidine it is best to first try other compound s that are thiol reactive. Other thio l reactive labels that have been used in previous studies include [Ru(bpy) 2 ( 5,6 epoxy 5,6 dihydro 1,10 phenanthroline)] 2+ [Ru(bpy) 2 ( 5 maleinimide 1,10 phenanthroline )] 2+ and [Ru(bpy) 2 ( 4 bromomethyl 4' methylbipyridine )] 2 + 17 21 22 Several experiments can be performed on a ruthenium labeled sample of K375C/C383A OxDC. Kinetics assays of Ru OxDC without the presence of oxygen but with a suitable quencher and 450 nm wavelength light ( compared against controls without light, without the ruthenium label, and with out the quencher) could confirm the viabi lity of LRET. Oxygen is a necessary cofactor, acting as a temporary electron sink. 23 If catalytic ability of Ru OxDC is retained in flash quench conditions with the absence of oxygen, it would suggest that the C terminal mangan ese is accepting electrons from the N terminal manganese.


25 Conclusion Theoretical calculations using the Marcus equation suggest that electron transfer between the ruthenium complex and C terminal manganese in Ru OxDC is possible, with a tunneling time of 660 n s However, further modification of the photosensitizer used may be needed to produce a sufficiently small tunneling time. A K375C/C383A OxDC mutant was successfully produced and can be used for future labeling at the C terminal manganese site. Th e mutant retained wild type kinetics, with K M = 10 2 mM and k cat = 90 13 s 1 MS experiments suggest that the ruthenium compound failed to bind to K375C/C383A OxDC, either at cysteine 375 or at a lysine or histidine. Future labeling should be attempted with a ligand containing an epoxide, malinimide, or bromo functional group Kinetic assays without the presence of oxygen should be performed on Ru OxDC in order to further investigate the viability of LRET.


26 Acknowledgments I would like to than k Dr. Alexander Angerhofer my thesis advisor, for his support and guidance during this project. I would also like to thank Anthony Pastore and indeed all the other members of the Angerhofer research group, for all of their help and advice. Finally I wou ld like to thank Dr. Kari Basso for her help with the mass spectrometry experiments. References 1. Twahir, U T. Investigations into the Enzymatic Mechanism of Bacillus Subtilis Oxalate Decarboxylase: An Electron Paramagnetic Resonance Approach. Ph.D. Disser tation, University of Florida, Gainesville, FL, 2015. 2. Richardson, K. E. and Tolbert, N. E. Oxidation of Glyoxylic Acid to Oxalic Acid by Glycolic Acid Oxidase. J. Biol. Chem 1961, 236 1280 1284 3. Pinzauti, G.; Giachetti, E.; Camici, G.; Manao, G.; Cappu gi, G.; and Vanni, P. An isocitrate lyase of higher plants: Analysis and comparison of some molecular properties. Arch. Biochem. Biophys. 1986 244 85 93. 4. Frassetto, L. ; Kohlstadt, I. Treatment and Prevention of Kidney Stones: An Update. Am. Fam. Physici an. 2011 84 1234 1242. 5. Rudie, A. W.; Hart, P. W. Mineral Scale Management Part II: Fundamental Chemistry. TAPPI J 2006 5 17 23. 6. Just, V. J.; Burrell, M. R.; Bowater, L.; McRobbie, I.; Stevenson, C. E. M.; and Lawson, D. The identity of the active si te of oxalate decarboxylase and the importance of the stability of active site lid conformations. Bornemann, S. Biochem. J 2007 407 397 406. 7. Cleland, W.W. ; Richards, N.G Investigating the roles of putative active site residues in the oxalate decarboxylase from Bacillus subtilis. Arch B iochem B iophy 2007 464 pp 36 47. 8. Pastore, T., u npublished. 9. Twahir, U. T.; Stedwell, C. N; Lee, C. T.; Richards, N. G. J.; Polfer, N. C.; Angerhofer, A. Free Radic. Biol. Med. 2015 80 59 66.


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