Site-Directed Mutagenesis and Ruthenium Labeling of Oxalate Decarboxylase

Timothy DeMason Undergraduate Honors Thesis – Spring 2018 Department of Chemistry, University of Florida

Contents List of Abbreviations ...... 3 Abstract ...... 4 Introduction ...... 5 Methods...... 11 Results and Discussion ...... 16 Calculation of Ru—Mn Electron Transfer ...... 16 Mutagenesis of K375C/C383A ...... 18 Synthesis of Ru-OxDC ...... 22 Conclusion ...... 25 Acknowledgements ...... 26 References ...... 26

List of Abbreviations bis-tris 2-bis(2-hydroxyethyl)amino-2-(hydroxymethyl)-1,3-propanediol bpy 2,2’-bipyridine

DMSO dimethyl sulfoxide

DTT dithiothreitol

E. coli Escherichia coli

FDH Formate Dehydrogenase

IA-phen 5-iodoacetamido-1,10-phenanthroline

IPTG isopropyl β-D-1-thiogalactopyranoside

LRET Long Range Electron Transfer mM mili Molar

μM micro Molar

MS Mass Spectrometry

NAD+ nicotinamide adenine dinucleotide

OxDC Oxalate Decarboxylase

PCET Proton Coupled Electron Transfer

PCR Polymerase Chain Reaction

PDB Protein Data Bank

Ru-OxDC Ruthenium Modified Oxalate Decarboxylase

Tris 2-amino-2-(hydroxymethyl)-1,3-propanediol

WT Wild Type Oxalate Decarboxylase

Abstract 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

2+ thiol reactive ruthenium compound [Ru(bpy)2(IA-phen)] . 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,

-1 with KM = 10 ± 2 mM and kcat = 90 ± 13 s . Attempts at labeling K375C/C383A OxDC with

2+ [Ru(bpy)2(IA-phen)] were unsuccessful. The failure of the labeling reaction appears to be due

2+ to an inability of [Ru(bpy)2(IA-phen)] to react with K375C/C383A OxDC rather than nonspecificity of the reaction. Future labeling with other thiol reactive ligands should be attempted.

Introduction Oxalic acid is a dicarboxylic acid with pKa’s of 1.2 and 4.2.1 Oxalic acid is produced in plants through several biochemical pathways including the activity of glyoxalate oxidase and isocitrate 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 relatively insoluble and is present in about

60% of kidney stones.4 Developing ways of reducing the degree of oxalate accumulation in the kidneys is of medical importance. The buildup of calcium oxalate deposits can also cause problems 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 degradation of oxalate into 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 Schemes 1A and 1B.1

– O O O O2 A CO2 + – O OH H O

– O O B + O H 2 CO H O + 2 + 2 + 2 2 O OH 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 1C).

The structure of the monomer is shown in Figures 1A and 1B and consists of two β-barrel domains, one at the N-terminal end (shown in green in Figures 1A and 1B) and another at the C- 6 terminal end (shown in blue in Figures 1A and 1B). Inside each β-barrel rests a manganese ion bound to three histidines and one glutamate, leaving two sites free for other small ligands.

A) C)

Figure 1: The crystal structure of Bacillus subtilis OxDC showing the manganese ions 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 PDB 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 dioxygen bind to the N-terminal manganese, with the dioxygen generating Mn3+.1 Then, glutamate-162 removes the remaining acidic proton from oxalate, while simultaneously an electron is transferred from oxalate to the manganese, i.e. proton coupled electron transfer

(PCET).1 Heterolytic cleavage of the oxalate 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 Mn2+ to 7 regenerate Mn3+. Concurrently, glutamate-162 protonates the carbon of the carbon dioxide radical 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

- peroxycarbonate (HCO4 ) which 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-terminal manganese binds oxalate (and oxygen as a co-catalyst) while it cycles between Mn2+ and Mn3+. From (9) with permission from Elsevier.

While the C-terminal manganese was originally thought to only play a structural role, further investigation into the OxDC mechanism suggests 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 ions is possible. Substitution of tryptophan-96 and tryptophan-274 with 8 phenylalanine or tyrosine leads to significantly reduced catalytic capability indicating their importance in catalysis.1,8 Additionally, EPR spin trapping studies of a flexible lid mutant suggest that the carbon dioxide radical anions and superoxide radicals are produced in separate locations.9 Since oxalate is known to bind at the N-terminal manganese, perhaps oxygen binds 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).1010

C-terminal N-terminal O O Glu Glu Glu Glu 280 101 280 101 – His + HC O - O O His 319 O 140 OH2 2 4 His 319 O His 140 2+ 2+ 2+ 2+ Mn Mn Mn Mn

His 275 OH2 His 97 OH2 His 275 OH2 His 97 O OH O His O His 273 His 95 His 273 95

– – Glu O Glu162 O 162 LRET + H+

OH OH Glu Glu Glu Glu 280 101 – 280 101 – O PCET O O His 319 O His 140 O His 319 O His 140 2+ 2+ oxidation 2+ 3+ Mn Mn + Mn Mn C – His 275 OH2 His 97 O O His 275 OH2 His 97 O OH His O His His 273 95 His 273 95 O

HO Glu162 – O Glu162

- - CO2 - CHOO - + HC2O4

OH OH Glu Glu Glu 280 101 PCET 280 Glu101 His His 319 O 140 reduction His 319 O His 140 2+ 2+ 2+ 3+ H Mn Mn – Mn Mn C His His – 275 OH2 97 O O His 275 OH2 His 97 O O His His His 273 95 O His 273 95 O

– HO Glu162 O Glu162 Figure 3: New proposed mechanism for OxDC, including the possibility of long range electron transfer, assuming oxalate binds in a bi-dentate fashion. From (10).

Selective oxidation of the C-terminal manganese could be used to further study the possibility of LRET. Ruthenium modified 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-called “flash-quench” technique shown in

Scheme 2A. A ruthenium(II) diimine complex, such as tris(bipyridine)ruthenium(II), is excited with a “flash” of light at 450 nm to produce an excited state which is then “quenched” by either an oxidant or reductant to produce ruthenium(III) or ruthenium(I), respectively.12 The oxidized or reduced ruthenium can then either remove or inject an electron from a suitable nearby target.12

2+ A) 2+ B) *Ru *Ru Q -0.8 V +0.8 V Q- hν hν

3+ 2+ 3+ 2+ + Ru Ru Ru Ru Ru +1.3 V -1.3 V 2+ 3+ Mn Mn Scheme 2: A) General flash-quench scheme showing both the oxidation and reduction routes with reduction potentials. B) Oxidation scheme proposed for OxDC using a small quencher, Q. Modified from (12) with permission under Caltech’s Open Access Policy.

In the case of OxDC, bis(bipyridine)(5-iodoacetamido-1,10-phenanthroline)ruthenium(II)

2+ (i.e. [Ru(bpy)2(IA-phen)] ) can be used to oxidize the C-terminal manganese (see Scheme 2B).

2+ The iodoacetamide group of [Ru(bpy)2(IA-phen)] is expected to react with reduced, surface

12 accessible cysteines though an SN2 mechanism as shown in Scheme 3. 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. 10

N N

N N N N 2+ O 2+ O NH Ru Ru I S O N N NH N N NH N N – NH S

2+ Scheme 3: Reaction of [Ru(bpy)2(IA-phen)] with a cysteine residue.

In order to label OxDC with this ruthenium 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 Angerhofer Lab. This mutant served as the starting place for this project, since the only cysteine in the OxDC sequence has been removed.

Methods Mutagenesis was performed using a Q5 Site-Directed Mutagenesis Kit from New

England Biolabs. Briefly, primers for two separate mutants were designed using NEBaseChanger to incorporate the cysteine TGC codon at the appropriate site (see Table 1) and purchased from

Integrated DNA Technologies.

Table 1: Site Directed Mutagenesis Primers

Primer Sequence (5’ to 3’) K375C FWD TTCAAAAGAAtgcCACCCAGTAGTGAAAAAG K375C 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

μM of both forward and reverse primers, and Q5 Hot Start High-Fidelity 1X 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 56, 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 10 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 competent E. coli cells were then transformed with 30 s of heat shock at 42 °C. The transformed cells were streaked onto LB agar plates treated with 50 ng/μL ampicillin and grown overnight. A colony was then selected and grown in an overnight culture for plasmid purification the following day. The miniprep was performed using 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) competent E. coli cells which were grown for a glycerol stock, 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 grown in an overnight culture using ampicillin-treated LB media (50 ng/μL 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 was grown in 3 L of ampicillin-treated LB media at 37 °C until OD600=0.5. At that point, the cells were heat shocked at 42 °C for 10 minutes and 5 mM MnCl2 and 0.8 mM isopropyl β-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 (50 mM Tris, 500 mM NaCl, 10 μM MnCl2, 10 mM imidazole, pH 7.5) before being sonicated. The lysed cells were pelleted down and the supernatant poured 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 Tris, 500 mM NaCl, 20 mM imidazole, 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). Fractions of every eluate were collected every 5-10 mL. These fractions were dialyzed overnight in 2 L of storage buffer (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 were 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.

O + FDH NAD CO + 2 + NADH H OH Scheme 4: Reduction of NAD+ by formate, catalyzed by FDH.

The assay was performed by mixing a 5 μL aliquot of OxDC with 99 μL of 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 between 2 and

100 mM. The samples were reacted for 1 minute at 25 °C before 10 μL of 1 M NaOH was added to quench the reaction. Then, 55 μL aliquots were mixed with 945 μL of 50 mM pH 7.8 phosphate buffer, 1.5 mM NAD+, and 0.0004% (m/v) FDH and incubated overnight at 37 °C.

Finally, absorbance readings 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 conditions. Both 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 sample at 4 °C for 2 hours using 2 L of pH 8 storage buffer. A solution of ruthenium label was prepared by dissolving between 1 and 2 mg of

[Ru(bpy)2(IA-phen)](PF6)2 (purchased from Santa Cruz Biotech) was in 1 mL DMSO and further diluted with 1 mL pH 8 storage 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 prepared 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, samples of protein 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 (NH4)HCO3. The gel band was then rehydrated with 12 ng/ml sequencing grade trypsin in 0.01% ProteaseMAX Surfactant and then overlaid with 40 µL of 0.01% ProteaseMAX

Surfactant and 50 mM (NH4)HCO3 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 UltiMate™ 3000

RSLCnano. The mobile phase A was 0.1% formic acid and acetic acid in water and the mobile phase B was acetonitrile with 0.1% formic acid in water. First, 5 μL of the sample was injected onto a 2 cm C18 column and washed with mobile phase A. The injector port was switched to 15 inject and the peptides were eluted off the trap onto a 25 cm C18 column for chromatographic separation. Peptides 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. The total run time was 60 minutes. The EASY Spray source operated with a spray voltage of 1.5 kV and a capillary temperature of 200oC. The scan sequence of the mass spectrometer was based on the TopTen™ method.

Results and Discussion

Calculation of Ru—Mn 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

13 of electron transfer, kET:

3 o 2 4휋 2 (Δ퐺 +휆) (1) 푘ET = √ 2 |퐻AB| exp (− ) ℎ 휆푘B푇 4휆푘B푇

where λ is the reorganization parameter, HAB is the electronic coupling between reactants and

11 products, and ΔG° is the reaction driving force. The electronic coupling factor, HAB, 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

2+ and the radius of the [Ru(bpy)3] complex. The Mn—S distance was estimated to be 11.1 Å from using the crystal structure OxDC (PDB 1UW8) to find the distance between the Mn and the

2+ 14 γ-carbon of lysine-375. The diameter of [Ru(bpy)3] was reported as 5.4 Å in the literature.

Figure 4: Plot of HAB vs donor-acceptor distance (RDA) for thermal (magenta) and optical (blue) intramolecular ET. From (11) with permission from the ACS. 17

Thus, the Mn—Ru distance was taken to be approximately 16.5 Å so, by using Figure 4 HAB is approximately 0.1 cm-1. The value of λ was estimated to be 0.8 eV in accordance the typical reorganization parameter for protein ET processes.11 The driving force of the reaction can be

2+ calculated using the reduction potential of [Ru(bpy)3] and the oxidation potential of a OxDC manganese model complex as estimates.15,16

Ru3+ + e- → Ru2+, E°= +1.3 V (vs NHE)

Mn2+ → Mn3+ + e-, E°= -0.73 V (vs NHE)

Taking ΔG° = -0.57 eV along with the other previous stated parameters and using Equation 1, we

6 -1 find that kET = 1.5 × 10 s with a tunneling time constant of τET = 660 ns. Because τET is on the order of hundreds of nanoseconds it is expected that the Mn—Ru electron transfer will occur, since electron transfer has been observed in similar ruthenium modified proteins with τET 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 changes in either of these parameters could have noticeable impacts on the rate of electron transfer. To minimize the tunneling time, it may be necessary to find a photosensitizer that produces a driving force of -ΔG° ≈ λ. The 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 d6 metal such as rhenium(I) or osmium(II) to further adjust the reduction potential.18

Mutagenesis of K375C/C383A

Based on the crystal structure of OxDC, alanine-341 and lysine-375 appeared to be the closest surface accessible residues to the C-terminal manganese with distances of 10.1 and 11.1

Å, 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 with distances between the residue and the manganese given too. This figure was generated in PyMOL using the 1UW8 PDB 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 seen 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

1 2 3 4 5 6 7 8 9 10

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.

Samples 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 sequence of the reverse strand did. Figure 7 shows the reverse complement of a section of the reverse strand sequencing data. This region displays the same sequence as the forward primer, indicating that the mutation was successfully incorporated into the synthesized 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. 20

Additionally, the sequencing data was compared to the sequence of OxDC (with the cysteine mutation included) using the NCBI Nucleotide BLAST. Both the forward and reverse sequence matched the comparison sequence for the bases sequenced.

The kinetics of the C383A and K375C/C383A mutants were 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.

0.16 0.14

0.12

) 1 - 0.1

0.08

(mg∙U 1

- 0.06 v 0.04 0.02 0 0 0.1 0.2 0.3 0.4 0.5 0.6 -1 -1 [oxalate] (mM ) Figure 8: Lineweaver-Burk plot constructed for the C383A mutant. A linear regression was performed generating a line given by y=0.142669x+0.00999 with R2=0.959.

0.07

0.06

0.05

) 1 - 0.04

(mg∙U 0.03

- v 0.02

0.01

0 0 0.1 0.2 0.3 0.4 0.5 0.6 [oxalate]-1 (mM-1)

Figure 9: Lineweaver-Burk plot constructed for the K375C/C383A mutant. A linear regression was performed generating a line given by y= 0.079825x+ 0.00802 with R2=0.983. 21

From these Lineweaver-Burk plots, values of KM and kcat 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

-1 KM (mM) kcat (s ) C383A 14 ± 4 73 ± 15 K375C/C383A 10 ± 2 90 ± 13 WT7 5 ± 1 28 ± 1

WT19 6.6 ± 0.6 71 ± 6 WT20 12 ± 3 158 ± 13

When accounting for the margin of error, the values for KM and kcat 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

2+ suitable candidate for labeling with [Ru(bpy)2(IA-phen)] .

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 described previously to identify if either the full ruthenium label or the phenanthroline ligand was bound to cysteine-375. The MS results did not 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 was not bound to the enzyme.

It was hypothesized that the cysteine may not be surface accessible at 4 °C, so an

Ellman’s assay was performed to determine the amount 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. Therefore, the labeling process was reperformed at 25 °C. The MS 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 yellow-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 expected, a potential structure of the Ru-OxDC protein was created in PyMOL, using the mutagenesis and bond fusing features to

2+ perform the K375C mutation and attach [Ru(bpy)2(IA-phen)] to the sulfur (Figure 10A). 23

A) B)

Figure 10: A) Estimated location of the [Ru(bpy)(IA-phen)]2+ complex relative to the C-terminal Mn. B) Residues within 6 Å displayed in orange. Figures generated in PyMOL using the 1UW8 PDB file.

After the structure was created, the residues within 6 Å were displayed to check for any steric interference. Figure 10B shows this display and reveals that there is sufficient space for the ruthenium compound to sit above the opening of the C-terminal β-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 determine 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 experiments.8 Again, the MS results did not show a labeled fragment.

Another possibility is that the unprotontated nitrogen of a histidine residue could react with the label’s iodoactamide group. However, this is not likely since histadine-376 is surface 24 accessible and is adjacent to cysteine-375 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-terminal 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 functional 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 compounds that are thiol reactive. Other thiol reactive labels that have been used in previous

2+ studies include [Ru(bpy)2(5,6-epoxy-5,6-dihydro-1,10-phenanthroline)] , [Ru(bpy)2(5‐

2+ maleinimide‐1,10‐phenanthroline)] , 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 without the quencher) could confirm the viability 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 manganese is accepting electrons from the N-terminal manganese.

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 ns. 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. The mutant

-1 retained wild type kinetics, with KM = 10 ± 2 mM and kcat = 90 ± 13 s .

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.

Acknowledgments I would like to thank 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 would 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. Dissertation, University of Florida, Gainesville, FL, 2015.

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