Many important biological processes involve electron transfer reactions. In the Stankovich group. We use the electron transfer reaction to learn more about various proteins. The electron transfer path is able to tell us about changes that occur in a protein when it binds to another protein/subunit or substrate. We are interested in the substrate molecules and their interactions with enzyme to form products We are interested in the synthesis of deoxyribonucleotides, which are formed through the reduction of ribonucleotides with the help of the enzyme ribonucleotide reductase. Ribonucleotide reductase is essential to normal cells. If the mechanism of ribonucleotide reductase could be fully understood, the knowledge could be used to restrict DNA synthesis, and therefore restrict the growth of tumors or virus replications (Sjöberg 1983). This is vitally important, because it could lead to information that could help find a cure for cancer.
Importance of the Stankovich Research
Ribonucleotide reductase plays an important role
in DNA biosynthesis, which makes the enzyme a favorable "target" for antiviral
and antitumor agents (Stubbe 1990). Recently, it was found that women
with breast cancer who used hormone receptors had a high concentration
of ribonucleotide reductase. Full characterization of ribonucleotide
reductase could allow the cancerous enzymes to be inhibited, while allowing
the "healthy" enzymes to work as normal. This could result in the
development of drugs which combat the many types of cancer that take millions
of lives each year.
Background
The enzyme ribonucleotide reductase (RNR) is involved
in a crucial step in DNA biosynthesis, the reduction of ribonucleotides
to deoxyribonucleotides. This reaction substitutes a hydrogen
for the 2' hydroxyl group of the ribose, with retention of configuration
to form the deoxyribonucleotide.
Ribonucleotide reductase is an ubiquitous protein
found in all living organisms, that can be divided into four distinct classes. Each class contains an essential organic radical, which
is formed through an oxidation-reduction reaction at the metal center (Stubbe
1990). Class I reductases, the mammalian form of RNR, are tetramers
composed of two proteins: R1 and R2. The protein can be purified from heat-inducible
overproducing strains of E.coli. (Sjöberg 1986). The X-ray crystal
structures of the R1 component (Uhlin 1994) and the R2 component have been
resolved (Nordlund 1993). Ribonucleotide diphosphate reductase from
aerobic E. coli is archetypal of these enzymes, which contain a diiron
cluster and a tyrosyl radical (Stubbe 1989). The sole member of Class
II ribonucleotide reductase is ribonucleotide triphosphate reductase from
Lactobacillus leichmanni. (Lammers 1983). This enzyme contains a dimanganese
cluster with a tyrosyl radical. Class III, again, has only one representative
enzyme. The enzyme isolated from Brevibacerium ammoniagenes, contains
a cobalt center with an adenosyl radical. A ribonucleotide enzyme
which can be isolated from E. coli, grown anaerobically, represents Class
IV (Ashley 1986). It contains an iron?sulfur center with a glycyl
radical. The Stankovich group’s research focuses on the Class I reductase.
Mechanism of Ribonucleotide Reductase
The proposed mechanism of substrate reduction (Figure
2) occurs at the substrate binding site of the enzyme (Lammers 1983).
A hydrogen is abstracted from the radical of the protein at the 3' carbon
of the substrate. The hydroxyl group at the 2' position gains a proton
and is then taken off as water. A 3' ketone then further reduces
the substrate, to form a 3'-deoxyribonucleotide radical concomitant, while
oxidizing the two cysteine thiols, which causes the formation of a disulfide
bond. From that point, the 3' carbon of the substrate gains a hydrogen
again, which forms the product (deoxyribonucleotide), which then will dissociate
from the active site. Once this has occurred, the reduced active
site must be regenerated, which means that the protein disulfide bond has
to be reduced through the transfer of reducing equivalents through an electron
transfer chain. This will occur by NADPH donating electrons that
will be transported via one of two flavoprotein reductase systems: thioredoxin/thioredoxin
reductase or glutaredoxin/glutathione reductase + glutathione (Sjöberg
1983).
R1
R1 catalyzes the reduction of substrate and contains
the binding sites for purines, pyrimidines, and allosteric cofactors (Mao
1992a). The X-ray crystal structure reveals that R1 is composed of three
domains: a helical N-terminal domain, a a/b barrel domain, and a abaab
region (Uhlin 1994). Cys225 and Cys462 lie on one side of the a/b
barrel domain in the crystal structure of the reduced protein. In
the crystal structure of the oxidized protein, the thiols from the residues
form a disulfide bridge, which led to the idea that the two residues supply
the reducing equivalents for substrate reduction by formation of a redox
pair, which was suggested beforehand via site-directed mutagenesis (Mao
1992b). The active site is located in a buried orifice between the
N-terminal and b-barrel regions. . R1 is composed of two polypeptide chains (a2), each
containing 761 residues, yielding a molecular weight of 171 kDa.
One hypothesis is that R1 is very unstable and quite flexible when not
bound to R2. This flexibility would allow it to easily bind to R2.
The R1 dimer would then become more stable and conformationally rigid when
it is joined to the top of the heart-shaped R2 dimer. Thus a compact
complex would be formed, while the active site remains accessible to substrate.
R2
R2 is a heart shaped dimer (Figure ???) with a molecular
weight of 87 kDa (Lynch 1989). It is composed of two identical polypeptide
chains (b2), each containing 375 residues, which interact at an angle to
form the dimer (Haskin 1996). R2 contains 1.2 equivalents (per dimer)
of the catalytically essential tyrosyl radical (oY122). It
also contains 3.5 equivalents of Fe(III), assembled as µ-oxo bridged
dinuclear clusters that are necessary for the generation and stability
of the tyrosyl radical. The formation of the tyrosyl
radical occurs due to the reaction of the diiron cluster(II) with oxygen
(Equation 1).
Fe(II)-Fe(II) + Y-OH + e- + O2 Æ Fe(III)-O-Fe(III) + Y-O* + OH- (1)
R2 may exist in any of three stable redox states. The first is the catalytically active R2, containing
oY122 and [Fe(III)Fe(III)]. The second state is R2met, which is composed
of [Fe(III)Fe(III)] and Y122. The third state, R2red, is fully reduced
[Fe(II)Fe(II)] and Y122 (Elgren 19265). Because of the known stochiometry
amount of radical/dimer, in wild type R2, 33% of the protein can be found
in the met form, while the remaining 67% is in the active form.
The reaction of RNR is under study. Thus far, the redox
chemistry of the dinuclear iron centers has proven to be an excellent probe
into the enzyme mechanism. Reduction of the tyrosyl radical
in the catalytically active R2 results in the formation of R2met (Fe(III)Fe(III),
Y122), which has an absorbance at 325 and 370 nm (from the ?-? transitions
of the diiron cluster), but no sharp peak at 410 nm, (due to the tyrosyl
radical). This reduction to R2red can be accomplished via two routes:
through chemical or electrochemical reduction (Lynch 1989). The second,
our method of choice, uses methyl viologen as an electron mediator .
Direct electron transfer to the protein from the electrodes is not possible
since the diiron center and tyrosyl radical are buried deep within the
protein structure. Therefore, methyl viologen is reduced at the electrode
surface and then transported to the protein surface where it reduces the
protein. R2met can be returned to the native protein by full reduction
followed by the addition of oxygen.
Coulometric and chemical titrations of R2ox with
methyl viologen have shown that during the addition of 2.5 reducing equivalents
to the oxidized form of R2, there is no observed build-up of reduced methylviologen
(MVred), indicating fast transfer of electrons from reduced methyl viologen
to the protein electron acceptor. At this point in the reaction,
the tyrosyl radical is fully reduced and 33% of the clusters are reduced
(Fe(II)Fe(II)), while 66% remain Fe(III)Fe(III). Further addition
of reducing equivalents results in the formation of MVred which is then
slowly oxidized by the remaining diferric clusters in R2. This suggests
that the rate of electron transfer from MVred to R2 has slowed significantly.
Therefore, researchers have concluded that two different populations of
iron exist in R2, the fast reducing diiron cluster ("F") and the slow reducing
diiron cluster ("S"). It was also noted that exposing the reduced
F cluster to oxygen did not generate a radical. Moreover, upon the
addition of oxygen, the amount of radical produced directly correlated
to the number of reduced S clusters. Thus, there appears to be two
different roles for the iron centers: generation of the tyrosyl radical
upon exposure to oxygen and donation of the extra electron (Miller 1999).
It has been reported that there is a ratio of 3.3
iron clusters per 1 tyrosyl radical. This negates the idea that there
should be one tyrosyl radical per 2 iron clusters (i.e. two tyrosyl radicals
per 4 iron clusters). If this is the case, where is the extra electron
coming from? If the tyrosyl radical can not donate the electron,
what will? We propose that one of the dinuclear iron centers reacts
with oxygen to form the tyrosyl radical and the other acts as the electron
donor.
Future Research of the Stankovich Group
Previous work in the Stankovich lab will provide
important building blocks for much of the proposed work. It has been
observed that when the R1-R2 complex is formed, the redox potential is
-226 mV, which is a considerable negative shift of over 100 mV potential
of the unbound R2. This indicates that R1 plays a role in electron
transfer occurring in R2. This project is designed to probe the reactivity
properties of R1 bound to R2.
F and S Cluster of R2
Research will first focus on studying the electrochemical
potential difference between the S and F cluster. The redox chemistry
of the dinuclear iron cluster will be continued to be used as a probe of
the enzyme mechanism. First, it must be determined if there is a
potential difference between the S and F cluster. The S cluster will
be investigated by measuring its potential as the protein is quickly reduced,
adding 2.5 reducing equivalents (the number of reducing equivalents associated
with the fast phase) to the protein. Several redox potential measurements
will be taken during the fast reduction. The slow phase will be probed
by fully reducing R2 and allowing it to reoxidize. As it oxidizes,
the potential will be measured until it reaches the fast phase (measuring
from 4.7 reducing equivalents to 2.5 equivalents). Additionally,
the effect of R1 binding on the potentials of the F and S clusters, as
well as the kinetics of their formation will be studied, using stop flow
measurements.
R1 bound to R2
Another project will look at ribonucleotide reductase
(R1 bound to R2). Ribonucleotide reductase is quite puzzling, due
to the necessity of both R1 and R2 to form the catalytically active protein.
The tyrosyl radical abstracts a hydrogen, then reduction takes place by
the thiols. The tyrosyl radical is located in R2, while
the thiols and substrate binding sites are found in R1.
Many questions can be posed. How do R1 and R2 interact to form the
catalytically active protein and the active site? Does a conformational
change have to occur for binding to take place? Answers to these
questions can be found using electrochemistry.
Our lab has looked at R2 alone to observe the kinetics
of reduction and redox potential involved. Fortunately, R2ox is spectrally
active, with the tyrosyl radical having an absorbance at 410 nm and the
diiron center having absorbances at both 325 and 370 nm. It is interesting
to note that the redox potential of the diiron center shifts to -226 mV
when bound to R1. Therefore, there must be some sort of conformational
change occurring near the diiron center when R2 is bound to R1. Surprisingly,
the spectral properties (EPR and Mössbauer) of the diiron center did
not change when R1 was bound to R2. This indicates that electrochemical
methods are the best way to understand what is happening when R1 binds
to R2. Therefore, experiments should be performed on R1 bound to
R2 to observe the interactions and the conformational change involved.
What causes the redox potential to change when R1
is bound to R2?
Our hypothesis is that rearrangement by the hydrogen bonds along the electron
pathway, upon binding of R1, could possibly cause the redox potential of
the iron centers of R2 to change. This hypothesis will be explored
by mutating residues along the electron transfer pathway (Tyr730, Tyr731,
Tyr356, Trp48, Asp237, and His118). Residues along the R1/R2 interface
(Glu350, Tyr356, Ser343, and Val353) will also be examined via site-directed
mutagenesis. All mutants have been made by Britt-Marie Sjöberg’s
group, and she has generously given us the clones. The redox properties
of the mutants will be characterized, and the effect of R1 bound to R2
on the redox potential will be explored.
Site-directed mutagenesis has been performed on R2 alone to try to elucidate
the electron pathway in R2.
Y356
Climent changed the R2 tyrosine 356 to an
alanine (Climent 1992). Mutating with alanine basically deletes the ligand
(changing from a hydroxyl group to a methyl group). This mutation
results in the mutant being inactive due to the electron transfer path
being blocked and the electron not being able to move along its usual path,
but still forms the normal amount of the catalytically essential tyrosyl
radical.
Trp48
Tryptophan 48 has also been converted to tyrosine,
phenyl alanine, and glutamine. Only the W48Y formed the catalytically
essential tyrosyl radical, but yet again, the enzyme was inactive (Parkin
1998). The hydroxyl group was able to donate an electron to make
the catalytically essential radical.
Asp237
Aspartic acid 237 was changed by an alanine (deleting
the ligand to the iron), asparagine (adding an amine group), and glutamic
acid (adding a methyl in the R group). With the glutamic acid mutant,
the tyrosyl radical is formed and low enzymatic activity is seen (~10%
of wild type) (Ekberg 1997). This change could be due to the aspartate
only being one methyl group shorter than the glutamate. It has been
reported that the longer glutamate could possibly still form the necessary
hydrogen bonds and still allow electron transfer (Persson 1997).
Equation 7 enables us to relate the shift in redox potential
of our redox active species (R2) to the binding constants of R1 ligands
with the oxidized and reduced protein.
Substrate and Activation Binding
Another question to be answered involves substrate
binding and activation binding. How does substrate and allosteric
effector binding affect the interactions between R1 and R2. Binding
will increase the rate of reaction between R1 and R2, but will it also
affect the redox potential? Measuring the effect of substrate and
allosteric effector binding on the potential of the R1-R2 complex will
enable this question to be answered. The allosteric effectors that
will be investigated are ATP and dATP.
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