site-directed mutagenesis and ruthenium labeling...
TRANSCRIPT
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......................................................................................................................................... 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
3
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
4
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
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.
5
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
OHO
O–
+O
O–
H
O2
H+
++
O
OHO
O–
O2
2 CO2 +
CO2
H2O
2
A
B
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.
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
A)
B)
C)
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
Mn2+
Glu280
His 273
His 275
His 319
OH2
O
O
O
O–
Glu162
Mn2+
Glu101
His 95
His 97
His 140
O
O–
OH
O
PCEToxidation
PCETreduction
- CO2
LRET
+ H+
C-terminal N-terminal
+ HC2O
4
-
- CHOO-
+ HC2O4-
O
O–
Glu162
Mn2+
Glu101
His 95
His 97
His 140
OH2
OH2
Mn2+
Glu280
His 273
His 275
His 319
OH2
O
O
Mn2+
Glu280
His 273
His 275
His 319
OH2
O
OH
O
O–
Glu162
Mn3+
Glu101
His 95
His 97
His 140
O
O–
OH
O
Mn2+
Glu280
His 273
His 275
His 319
OH2
O
OH
O
OH Glu162
Mn2+
Glu101
His 95
His 97
His 140
O
O–
C+
O–
O
Mn2+
Glu101
His 95
His 97
His 140
OC–
O
O
OH Glu162
Mn2+
Glu280
His 273
His 275
His 319
OH2
O
OH
Mn3+
Glu101
His 95
His 97
His 140
O–
O
H
O
O–
Glu162
Mn2+
Glu280
His 273
His 275
His 319
OH2
O
OH
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).
9
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
Ru3+
Ru2+
Ru2+
Ru+
*
hν
-0.8 V
+1.3 V -1.3 V
+0.8 V
Ru3+
Ru2+
Ru2+*
Q
Q-
hν
Mn2+
Mn3+
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)
(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(bpy)2(IA-phen)]2+ is expected to react with reduced, surface
accessible cysteines though an SN2 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
Ru2+
N
N
N
N
N
N NHI
O
O
NHS–
Ru2+
N
N
N
N
N
N NHS
O
O
NH
Scheme 3: Reaction of [Ru(bpy)2(IA-phen)]2+ 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.
11
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
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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
H OH+ NAD+
FDHCO
2 NADH+
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.
16
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
of electron transfer, kET:13
(1) 𝑘ET = √4𝜋3
ℎ2𝜆𝑘B𝑇|𝐻AB|
2exp (−(Δ𝐺o+𝜆)2
4𝜆𝑘B𝑇)
where λ is the reorganization parameter, HAB is the electronic coupling between reactants and
products, and ΔG° is the reaction driving force.11 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
and the radius of the [Ru(bpy)3]2+ 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
γ-carbon of lysine-375. The diameter of [Ru(bpy)3]2+ was reported as 5.4 Å in the literature.14
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
calculated using the reduction potential of [Ru(bpy)3]2+ 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
find that kET = 1.5 × 106 s-1 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
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
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.
1 2 3 4 5 6 7 8 9 10
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.
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.
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.
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(m
g∙U
-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(m
g∙U
-1)
[oxalate]-1 (mM-1)
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
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
suitable candidate for labeling with [Ru(bpy)2(IA-phen)]2+.
KM (mM) kcat (s
-1)
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
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 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
perform the K375C mutation and attach [Ru(bpy)2(IA-phen)]2+ to the sulfur (Figure 10A).
23
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
A) B)
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
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 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.
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 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
retained wild type kinetics, with KM = 10 ± 2 mM and kcat = 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 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.
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