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132
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THS

    

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THESIS

lllllllllllllllllllllllllUllllHillllllllllllllllll

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thesis entitled

DNA Photolyase: An Electron
Paramagnetic Resonance Spectroscopy Study

presented by

Kristi Lee Westphal

has been accepted towards fulfillment
of the requirements for

Masters Chemistry

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1/98 C'JCIHCIDmDUOpGS‘QM

DNA PHOTOLYASE: AN ELECTRON PARAMAGNETIC
RESONANCE SPECTROSCOPY STUDY

By

Kristi Lee Westphal

A THESIS
Submitted to
Michigan State University
in partial fulfillment of the requirements

for the degree of

MASTER OF SCIENCE

Department of Chemistry

1997

ABSTRACT

DNA PHOTOLYASE: AN ELECTRON PARAMAGNETIC
RESONANCE SPECTROSCOPY STUDY

By

Kristi Lee Westphal

DNA photolyase is an enzyme that repairs thymine dimers formed by exposure of
the DNA strand to far UV light. This enzyme is activated when it is in the presence of
visible light. The basic mechanism of photolyase is as follows: (1) photolyase binds to
the dimer in a light independent step (2) upon absorption of a photon the dimer is split (3)
photolyase dissociates from the DNA. Photolyase has been successfully overproduced in
E. coli cells and can now be isolated in a stable, nearly pure form.

Electron Paramagnetic Resonance (EPR) experiments have been conducted on
photolyase in order to better understand the repair mechanism. A spin polarized EPR
signal has been detected using time resolved spectroscopy. Labeling techniques have
been used in order to identify the active species involved in the spin polarization process.

Experiments which have been performed and future work in this area will be discussed.

ACKNOWLEDGMENTS

I would like to thank my colleagues for all of their support, technical and
personal. I want to give a huge thank you to Yvonne Gindt who is a great teacher and has
taught me all that I know on this research. I would also like to thank my advisor, Jerry
Babcock, for his guidance and for teaching me the difference between ‘which’ and ‘that’!
A special thanks goes out to my mother whose words of encouragement always keeps me

going when the only thing I want to do is give up. Thanks to all of you!

iii

TABLE OF CONTENTS

LIST OF FIGURES ............................................................................................................ v

ABBREVIATIONS ................................................................................ vii

1. CHAPTER 1: INTRODUCTION .......................................................................... 1
11. CHAPTER 2: ISOLATION AND CHARACTERIZATION

OF PHOTOLYASE ...................................................................... 25

A. Materials and Methods .............................................................................. 26

1. Bacterial Strains and Plasmids ...................................................... 26

2. Chemicals ...................................................................................... 26

3. Buffers ........................................................................................... 26

4. Equipment ..................................................................................... 27

5. Assay of DNA Photolyase ............................................................ 27

B. Results ....................................................................................................... 27

1. Cell Growth ................................................................................... 27

2. Isolation ......................................................................................... 28

3. Activity of the Purified Enzyme ................................................... 36

C. Discussion ................................................................................................. 39

D. References ................................................................................................. 40

III.

IV.

CHAPTER 3: IDENTIFICATION OF THE TRANSIENT, SPIN

A.
B.
C.

POLARIZED EPR SIGNAL OF PHOTOLYASE ....................... 41
Materials and Methods .............................................................................. 44
Results ....................................................................................................... 45
References ................................................................................................. 66

CHAPTER 4: ISOTOPIC LABELING STUDIES PERFORMED

POP”?

PHOTOLYASE ............................................................................ 67
Materials and Methods .............................................................................. 71
Results ....................................................................................................... 73
Future Work .............................................................................................. 80
References ................................................................................................. 82

iv

Figure 1.1.

Figure 1.2.
Figure 1.3.

Figure 1.4.

Figure 1.5.
Figure 2.1.
Figure 2.2.
Figure 2.3.
Figure 2.4.
Figure 3.1.

Figure 3.2.

Figure 3.3.

Figure 3.4.

Figure 3.5.

Figure 3.6.

LIST OF FIGURES

The four isomers of pyrimidine dimers ...................................................... 3
Basic mechanism of dimer splitting by photolyase .................................... 6
Chromophores of DNA photolyase ........................................................... 8
Absorption spectra of the oxidized, semiquinone,

and reduced oxidation states of DNA photolyase ..................................... 12
Proposed mechanism for dimer repair ...................................................... 16
Flow chart for procedure for isolating photolyase .................................... 31
SDS-PAGE gel of 95% pure photolyase .................................................. 33
Absorption spectrum of photolyase in its semiquinone form ..................... 35
Activity assay of photolyase in HEPES buffer and in Tris buffer ................ 38
Activation of photolyase by electron transfer from trp-306 ....................... 43
Steady state spectrum of photolyase in its semiquinone form at

a concentration of 62.5 uM ........................................................................ 47
Transient EPR spectrum of photolyase in its oxidized form at a

concentration of 73.5 11M ......................................................................... 49
Kinetic trace of the transient radical with photolyase in its
oxidized state at a concentration of 84.8 uM ............................................ 51
Proposed reaction mechanism for photoreduction that includes

the spin polarized species thought to give the transient EPR signal ............. 54
Absorption spectrum of photolyase in its semiquinone form ..................... 56

Figure 3.7. Absorption spectrum of photolyase in its oxidized form ........................... 60

Figure 3.8. Amplitude of transient signal versus
amplitude of steady state signal ...................................................................... 62

Figure 3.9. Proposed reaction mechanism with the fully oxidized flavin
as a member of the spin polarized pair .................................................... 64

Figure 4.1. Numbering scheme for the tryptophan molecule ..................................... 70

Figure 4.2. Absorption spectrum of the enzyme grown on M-9
medium with unlabeled tryptophan .......................................................... 75

Figure 4.3. Kinetic trace of the transient radical obtained with the
control enzyme sample in its oxidized state at a concentration
of 0.133 mM and 300 flashes averaged .................................................... 77

Figure 4.4. Kinetic trace of the transient radical obtained with the

control enzyme sample in its oxidized state at a
concentration of 0.133 mM and 500 flashes averaged ................................. 79

vi

 

Abbreviations: pyr<>pyr, pyrimidine dimer; FADH, flavin adenine dinucleotide;
MTHF, methenyltetrahydrofolate; E-FAD, enzyme apoenzyme bound to flavin; T<>T,
thymine dimer; pyr, pyrimidine molecule; pyr", pyrmidine charged radical; tac, promoter
for transcription; phr, gene; NpT<>TprNp, DNA strand containing three of any bases
and two thymine bases; T<c,s>T, cis,syn cyclobutane dimer; trp, tryptophan molecule;

HOMO, highest occupied molecular orbital; LUMO, lowest unoccupied molecular orbital

vii

 

Chapter 1: Introduction

 

DNA photolyase is an enzyme that repairs cyclopyrirnidine dimers found in DNA.
These dimers are formed by adjacent bases on the same strand when the DNA has been
exposed to far UV light (200-300nm). The two main bases that are affected are purine
and pyrimidine that absorb at 9» max = 250-270nm, with pyrimidine (pyr, see page 25 for
abbreviations) photoproducts predominating.‘ Four isomers of pyrimidine dimers are
shown in Figure 1.1, with the cis, syn isomer being the natural substrate of photolyase.
The pyr<>pyr kills cells by blocking replication and transcription and by causing a
mutation at the site of the lesion. Cells protect themselves from these effects by
eliminating photodimers from their genome either by excision repair or by
photoreactivation. The latter process is carried out by DNA photolyase, which prevents
the harmful effects (cancer, mutations, death) of far UV light by concurrent or subsequent
exposure to near UV light (3 50-500nm).2 The DNA photolyases are a group of enzymes
that catalyze this photoreactivation of damaged cells by breaking the cyclobutane bonds
of the pyrimidine dimer, thereby restoring the pyrimidine monomers. These light-
activated repair enzymes are widespread in nature and have been found in many bacteria,
blue green algae, fungi, higher plants, and all major groups of vertebrates with the
possible exception of placental mammals. Its presence in humans has been a

controversial issue with the most recent conclusion being that photolyase is found in

humans.3

 

Figure 1.1. The four isomers of pyrimidine dimers.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

o 0
CH3 CH3 H H $3943 H
H 2 a s
\N 5: :5. N/ \N —5- N/
o/‘\ 6 6. X o/l\ 6 6' /J\o
N E a N o i A E T
F1 F1
J, ,L H H
cis-syn trans-syn (racemic)
o o o o
CH3|:| H 9H3? H
H\N ii is, N/ \N i 5. N/
0% 6 6' /Ko 02\ 6 6' /J\
N z"- N N l l N O
I LH3FI I I i" L“3|
H H H
cis-anti

trans-anti (racemic)

Heelis, P.F., Hartman, R.F., and Rose, S. Chemical Society
Reviews 1995, 24, 290. '

Figure 1.1

4

Photoreactivation was discovered in the late 1940’s by Albert Kelner when he
showed that after UV inactivation Streptomyces Griseus cells could be reactivated by
subsequent irradiation with visible light.4 In 1962, Rupert, et al. showed that
photoreactivation restores pyrimidine dimers to the original pyrimidines in DNA.5 By
1960 the basic mechanism was thought to occur in three steps: (1) Photolyase binds to
pyrimidine dimers in DNA in a light independent step (2) upon absorption of a photon
the cyclobutane dimers are split to yield two pyrimidines (3) photolyase then dissociates
from the DNA. Figure 1.2 shows a diagram of this basic mechanism.

The photolyase gene was cloned by Sancar and Rupert in 1978.6 In 1983 Sancar,
et al. described the construction of a t'ac-phr plasmid that greatly over produces
photolyase in Escherichia coli.7 These achievements in the cloning and overproducing of
photolyase made it possible to study this enzyme in considerable detail. Now with a
more efficient method for purification (described in chapter 2) substantial quantities of
the enzyme can be produced and isolated in a relatively short period of time.

DNA photolyases are monomeric proteins of molecular weight 55-65 kDa. Their
action is described by classic Michaelis-Menten enzyme kinetics with the exception that
the catalysis is light initiated. All contain stoichiometric amounts of two light absorbing
cofactors, one of which is 1,5-dihydroflavin adenine dinucleotide (FADH) and the other
either methenyltetrahydrofolate or 8-hydroxy-5-deazariboflavin.8 The structures of the
two Chromophores are shown in Figure 1.3. The folate class includes enzymes from
Escherichia coli, Neurospora crassa, Bucillusfirmus, and Saccharomyces cerevisiae, and

the deazaflavin class includes photolyases from Anacystis m'dulans, Streptomyces griseus,

Figure 1.2. Basic mechanism of dimer splitting by photolyase. PRE is the
photoreactivating enzyme, and pyr<>pyr is a pyrimidine dimer in
damaged DNA.

6

PRE + pyr<>pyr

k1

 

V

A

 

k2

PRE — pyr<>pyr

k3

 

V

hv

PRE + pyr + pyr

Figure 1.2

 

Figure 1.3. Chromophores of DNA photolyase.

 

8
R -
H3C / ILI N Y0
.1 NH

a)FADH'

O

i}
/ \ H
N HEE—coz’

Hofli/ (g

\N

/J\ 6202'
H2 N N N

A
b) 5, 10-MTHF
R
HO N N YO
N H
C) 8-HDF

Kim, S. and Sancar, A. Photochemistry and Photobiology 1993, 5 7, 896.

Figure 1.3

9

Scenedesmus acutus, Halobacterium halobium, and Methanobacterium
thermoautotrophicum.

In photolyase, the flavin chromophore can by found in all three of its oxidation
states: the fully oxidized FADOX, the semiquinone FADH', and the fully reduced FADH2
(FADH‘). The E-FADox form is catalytically inert9 and the E-FADH‘ form is active, but
with low quantum yield. The FADH‘ form is the active form of the enzyme with a
quantum yield between 0.5 and 0.9.lo Flavin is required for catalysis because it is the
photoactive species in the enzyme. It may also play a structural role, however, as the
flavin cofactor may provide extra stabilization to the enzyme by keeping the two helix
clusters together; the clusters probably have to move apart for binding of FAD to the
apoenzyme.”

In E. coli the second chromophore, MTHF, functions as a light harvesting
molecule. It absorbs a photon of light and transfers this energy to FADH. It is not
necessary for substrate binding and does not have to be present for dimer cleavage to
occur. The MTHF cofactor is located on the periphery of the enzyme. This explains why
enzyme lacking this cofactor retains its natural conformation and thymine dimer binding
affinity. Moreover, since the MTHF cofactor is solvent accessible, it readily dissociates
from the enzyme during isolation.

Why does photolyase need two chromophores when flavin is able to absorb light
directly and can donate an electron to the dimer? Flavins are ubiquitous as redox-active
components of many enzymes. Although flavins can absorb light, they are not very

efficient (e = 5,000 M"cm’1 at 350nm for FADH'). The folate is much more efficient at

 

10
this task (8 = 25,000 M"cm" at 390nm).12 The folate is thus used in an antenna function

to absorb light and to transfer it to the flavin, which is used for chemistry. Therefore
both chromophores are needed for efficient dimer repair.

The native form of the enzyme contains one mole of MTHF and one mole of
FADH per mole of apoenzyme. Its absorption spectrum is characterized by a peak at
280nm due to aromatic amino acid residues and a peak at 384nm due to enzyme bound
MTHF. All three oxidation states of the flavin cofactor in the enzyme have characteristic
absorption spectra,13 as can be seen in Figure 1.4. The semiquinone, or one electron
reduced form of the enzyme, is blue in color and the enzyme readily turns blue during
purification due to oxidation of flavin to the semiquinone form. The absorption spectrum
of the semiquinone form has peaks at 480, 580, and 625nm typical of the neutral flavin
radical. The oxidized form is bright yellow in color due to its high extinction coefficient
at its absorption maxima of 450nm (e = 11,300 M'1 cm"). Upon full reduction, the
enzyme turns a pale yellow and absorbs at 366nm (e = 5680 M'1 cm“).

In 1995, the crystal structure of photolyase was reported by Park et al. The
enzyme is made up of a single polypeptide chain of 471 amino acids and the light
harvesting and catalytic chromophores, both of which are noncovalently attached
chromophores. The polypeptide chain is folded into five [3 strands, 20 on helices, five
short 310 helices, and connecting segments. There are two major domains, an a/B domain
which contains the MTHF chromophore, and the helical domain that binds the FADH
chromophore. The two chromophores are separated by 16.8 A; the excitation energy is

transferred from MTHF to FADH' with a time constant of about 200ps and 62%

11

Figure 1.4. Absorption spectra of the oxidized, semiquinone, and reduced
oxidation states of DNA photolyase.

Extinction Coefficient (mM"cm")

' 6.0~

 

3.0"

 

300

 

 
   

1 1

400 500 600

Wavelength (nm)

Jorns, M.S., Wang, 8., Jordan, SP, and Chanderkar, L.P.,

Biochemistry 1990, 29, 560.

Figure 1.4

1 - Fully Oxidized
2 - Semiquinone
3 — Fully Reduced

700

 

13

efficiency.‘4

Substrate binding depends on both pyr<>pyr structure and the local structure
about the dimer in the DNA backbone with pyr<>pyr binding to the flat surface of the
enzyme’s helical domain. The enzyme binds a T<>T-containing substrate by contacting
the first phosphate 3’ to T<>T on the damaged strand and the phosphate opposite the
T<>T across the minor groove on the undamaged strand of a duplex. There is a
depression or “hole” in the helical domain of the enzyme that leads directly to the FAD
binding site. This hole has the right dimension to allow a thymine dimer to fit precisely
and come within van der Waals contact distance of the FAD. A thymine dimer has
asymmetric polarity, which fits well with the asymmetry of the hole as the residues lining
the hole are hydrophobic on one side and polar on the other. Binding in this fashion
would allow electron transfer between FAD and the substrate to occur over a short
distance and with high efficiency. The pyr<>pyr is proposed to bind in this way by
flipping out of the helix and into the hole.

Photolyase is a “structure specific” protein. This means that its catalytic
specificity is determined by the overall structure of the DNA in contrast to a “sequence
specific” protein, which relies on the sequence of the DNA for binding. The natural
substrate for the enzyme is a pyr<>pyr duplex. A minimum length of DNA is required
and the substrate must be at least a 5-mer of the form NpT<>TprNp.15 Most of the
binding free energy comes from the DNA backbone where the enzyme makes close
contact with the pentose part of the backbone. The base composition of the dimer is also

important with the T<>T composition giving the highest affinity for binding. The

14
secondary structure of the DNA strand also plays a role in binding. Photolyase can bind

to either UV-irradiated single- or double-stranded DNA with similar affinity.16 The
tertiary structure has no effect on activity and photolyase can repair relaxed and
supercoiled DNA with equal efficiency. The T<c,s>T is the natural substrate for the
enzyme.

The proposed repair mechanism of pyrimidine dimers formed in DNA is shown in
Figure 1.5. Photolyase binds to the dimer formed in the DNA strand independent of
light. The folate chromophore absorbs a 350-450nm photon and transfers the excitation
energy to the flavin chromophore. The photoexcited flavin then transfers an electron to
pyr<>pyr and the 5-5 and 6-6 bonds of the cyclobutane ring are broken leaving a pyr and
a pyr". The pyr" donates an electron back to the one-electron oxidized flavin
chromophore to regenerate FADH‘ along with two neutral pyrimidine molecules. The
enzyme then dissociates from the DNA fully intact and able to repair another dimer.” ‘8' '9

In addition to the light-induced electron transfer process that underlies dimer
repair, a second photoinduced electron transfer can be observed. When DNA photolyase
is isolated, one electron oxidation of the fully reduced active form of the enzyme may
occur. This semiquinone form is dark stable and catalytically inactive. Studies have
shown that in vitro photoreactivation proceeds by photoreduction of the flavin radical
before dimer repair can take place.2°' 2‘ Excitation of the flavin radical results in
intramolecular electron or hydrogen atom transfer from an amino acid residue to the
radical. Heelis, et al. studied the exited-state properties of photolyase in the semiquinone

form to investigate the identity of this amino acid.22 They acquired a flash-induced

15

Figure 1.5. Proposed mechanism for dimer repair.

 

e-

A

IFADH' + Dimer

i

hv

 

FADH- + Dimer

REACTANTS

k1=6x109S'1
——>

16

  

k3=109-10‘°s"

 

1

FADH' + Dimer"

k2=106S‘l (non-enzymatic rate)
or k2=109s1 (enzymatic rate)

A

FADH' + monomer"
+ monomer

PRODUCTS

Heelis, P.F., Hartman, R.F., and Rose, S.D., Chemical Society
Reviews 1995, 24, 295.

Figure 1.5

17
difference spectrum of photolyase and compared it to spectra of the only three amino

acids with absorptions above 350nm: tryptophan, tyrosine, and histidine. Only the
addition of the tryptophan spectrum to the enzyme spectrum coincided with the spectrum
observed following laser excitation of the enzyme. These results suggested that
tryptophan is the amino acid donor in the photolyase enzyme system.

Li et al. carried these results one step further to determine the specific tryptophan
in the enzyme that was responsible for donating the electron or hydrogen atom.” This
study involved the use of site-directed mutagenesis to replace all fifteen tryptophan
residues individually by phenylalanine. Using flash photolysis they found the trp-306
mutation caused photoreduction of FADH' to cease without effecting the excited state
properties of F ADH' or the substrate binding. This led to the conclusion that trp-306 was
the photoreductant. Kim et al. confirmed this assignment by the use of time-resolved
EPR studies.24 They obtained a transient EPR spectrum of Enz-FADH' induced by a flash
of light. The spectrum showed emissive and absorptive patterns, typical of spin-polarized
states, consisting of three components split by about 15G. Superimposed on each of the
three components is partially resolved fine structure with splittings of about 5G.
Photolyase grown in poor medium containing isotopically labeled tryptophan was used
to test the identity of the amino acid donor. Upon deuteration the fine structure collapses,
but the three major components are still observable, suggesting that the radical did arise
from a tryptophan molecule. To identify the particular tryptophan in photolyase that
photoreduces the flavin, site directed mutagenesis was performed, as above, with the

exception that EPR was used for analysis. The transient spectrum disappeared when trp-

18
306 was mutated, while for all other tryptophans no change in the transient could be

observed. From the results of these studies, the conclusion was made that the donor was
indeed a tryptophan molecule and further that it was trp-306.

These experiments led to the proposal that trp-306 is the amino acid donor in the
reactivation process that reduces the flavin semiquinone to its active fully reduced form.
In this mechanism, the trp-306 can either donate an electron or a hydrogen atom to the
photoexcited flavin, thereby producing the active form of the enzyme. These two
possibilities raise the issue of the ionization state of the reduced flavin cofactor in the
active enzyme and a number of studies have been carried out to determine whether the
neutral or anionic flavin hydroquinone is physiologically the more important. Hartman
and Rose studied reduced flavin model compounds and concluded that the anionic, single
protonated form is more active in dimer repair than the doubly protonated form.25
Photolysis of 1.2x10'5M flavin and 3.4mM N(l), N(3)—dimethyluracil cis-syn-cyclobutane
dimer in solutions ranging from pH 5.5 to 9.1 was carried out. A titration curve of the
splitting efficiency, centered at pH 7.7, clearly showed the flavin anion was
approximately eight times more active in dimer repair than the neutral form. In another
study, Ghisla et al. showed that the absorption spectra of reduced flavins are influenced
by the degree of planarity of the flavin molecule.26 Neutral reduced flavins posses
maximum planarity with a typical A,n,,~390nm. Ionization at N1 allows flexing along
the N5-N10 axis which results in a blue shift to 350i10nm. The reduced flavin in the
active form of photolyase has XW=366nm27 which may correspond to this flexing upon

ionization.

 

l9

Isotopic labeling is a third route used in identifying whether the donated species is
an electron or a hydrogen atom. Isotopic labeling studies have been conducted on the
tryptophan responsible for the donation of the electron or hydrogen atom. The neutral
and cationic forms of the tryptophan radical differ in unpaired electron spin densities. The
tryptophan cation radical has high spin density at the C(2) a-proton and at the C(3) B-
methylene carbon position while the neutral tryptophan radical has significant spin
density at the N(l) nitrogen and C(3) B-methylene carbon position.28 Therefore the two
forms can be discriminated by EPR spectroscopy when the tryptophan has been
isotopically labeled at these positions. A sample was specifically labeled at the a-ring
proton positions causing the smaller hyperfine couplings (~5G) to disappear. These
results suggest that the transient EPR spectrum arises from a cation rather than a neutral
tryptophan radical. Although these labeling studies were conducted on the FADox-trp
radical pair, the results may be carried over to the FADH‘ -trp system because the
tryptophan radical, not the flavin, of the radical pair is believed to give the transient EPR
signal.29 The observation that the active reduced flavin is present in its anionic form in
photolyase coincides with the conclusions that most photoexcited flavins reacting with a
tryptophan molecule are primarily electron transfer types of reactions.30

Now that an electron rather than a hydrogen atom transfer mechanism has been
proposed for photo-repair a question arises as to the direction of the electron transfer,
from reduced flavin to dimer or the reverse? When FADH‘ absorbs a photon, an electron
is promoted from the HOMO (it-orbital) to the LUMO (1t' orbital). Therefore lFADH'

becomes a good electron donor and an electron can be transferred from the n* orbital of

20
‘FADH‘ to the LUMO of the dimer. The reverse case, electron transfer from dimer to

FADH‘, would be unlikely because reduced flavins are much better donors than
acceptors.’ ‘

Another way to determine the direction of electron transfer is to calculate the free
energy change for both reactions. The equation used is as follows:

AG(kJ mol") = 96.5 [E° (D‘+ D) - E° (A A") - 0.026] - E0“,
where E° (D'+/D) and E°(A/A") represent the reduction potentials of the donor and
acceptor, and E0,O is the energy of the excited state of the flavin chromophore. The free
energy change for electron transfer from flavin to dimer is calculated to be AG = -125 kJ
mol'1 and from dimer to flavin is AG = +180 kJ mol". Therefore electron transfer from
flavin to dimer is energetically more favorable”.

The purpose of the project described in this thesis is investigation of the spin
polarized EPR signal of photolyase and the mechanism of electron transfer in dimer
repair. A new isolation procedure facilitated this research by providing a more stable
sample with which to work. This improved procedure has cut the time needed for
isolation of the enzyme by over two-thirds. In addition, a new buffer system has been
employed to improve the stability of the enzyme. Due to shorter preparation time and
increased stabilization, one can consistently expect to get a purified product in the one
electron reduced semiquinone form. Consistency of the end product and optical
spectroscopy allows for accurate characterization of the samples being used for

experimentation.

20

‘FADH' to the LUMO of the dimer. The reverse case, electron transfer from dimer to
FADH', would be unlikely because reduced flavins are much better donors than
acceptors.31

Another way to determine the direction of electron transfer is to calculate the free
energy change for both reactions. The equation used is as follows:

AG(kJ mol") = 96.5 [E° (D'+ D) - E° (A A") - 0.026] — E0,o
where E° (Di/D) and E°(A/A") represent the reduction potentials of the donor and
acceptor, and E,o is the energy of the excited state of the flavin chromophore. The free
energy change for electron transfer from flavin to dimer is calculated to be AG = -125 kJ
mol‘l and from dimer to flavin is AG = +180 kJ mol“. Therefore electron transfer from
flavin to dimer is energetically more favorable”.

The purpose of the project described in this thesis is investigation of the spin
polarized EPR signal of photolyase and the mechanism of electron transfer in dimer
repair. A new isolation procedure facilitated this research by providing a more stable
sample with which to work. This improved procedure has cut the time needed for
isolation of the enzyme by over two-thirds. In addition, a new buffer system has been
employed to improve the stability of the enzyme. Due to shorter preparation time and
increased stabilization, one can consistently expect to get a purified product in the one
electron reduced semiquinone form. Consistency of the end product and optical
spectroscopy allows for accurate characterization of the samples being used for

experimentation.

21

The identification of the spin polarized species giving the transient signal has also
been investigated by using deuterium labeled samples and EPR spectroscopy. The spin
polarized EPR signal observed may originate from either or both of the species making
up the spin polarized pair. Using labeled tryptophan species, we have observed that only
the tip radical contributes to the transient EPR spectrum as shown in chapter 4. These
results can contribute to the better understanding of this photobiochemical reaction that
perserves the integrity of the DNA strand. The dimer repair process performed by
photolyase is becoming especially crucial as the result of the increased incidence of UV

light reaching the earth’s surface over the past several years.

 

22

 

References

1. Wang, S.Y. Photochem. Photobiol. of Nucleic Acids 1976, Vols. 1, 11, Academic Press,

10.
ll.
12.
13.
14.
15.
16.
17.

18.

19.

NY.
Sancar, A. and Sancar, G.B. Annu. Rev. Biochem. 1988, 57, 29.

Sutherland, B.M. and Bennett, P.V. Proc. Natl. Acad. Sci. USA 1995, 92, 9732-9735.

. Kelner, A. Proc. Natl. Acad. Sci. USA 1949, 35, 73.

Wolf, D. And Rupert, C.S. Biophys. Biochem. Res. Commun. 1962, 7, 237.

Sancar, A. and Rupert, C.S. Gene 1978, 4, 294.

Sancar, G.B., Smith, F.W., and Sancar, A. Nucleic Acids Res. 1983, 11, 6667-6678.
Kim, S. and Sancar, A. Photochem. Photobiol., 1993, 5 7, 895.

Payne, G., Wills, M., Walsh, C., and Sancar, A. Biochemistry 1990, 29, 5706-5711.
Sancar, A. Biochemistry 1994, 33, 2-9.

Park, H., Kim, S.T., Sancar, A., and Deisenhofer, .1. Science 1995, 268, 1866-1872.
Heelis, P.F., Hartman, R.F., and Rose, S. Chem. Soc. Rev. 1995, 24, 289-297.
Jorns, M.S., Sancar, GB, and Sancar, A. Biochemistry 1984, 23, 2673-2679.
Park, H., Kim, S.T., Sancar, A., and Deisenhofer, J. Science 1995, 268, 1866-1872.
Jorns, M.S., Sancar, GB, and Sancar, A. Biochemistry 1985, 24, 1856-1861.
Sancar, G.B., Smith, F .W., and Sancar, A. Biochemistry 1985, 24, 1849-1855.
Heelis, P.F., Okamura, T., and Sancar, A. Biochemistry 1990, 29, 5694.

Kim, S.T., Sancar, A., Essenmacher, CE, and Babcock, G.T. J. Am. Chem. Soc.
1992, 114, 4442.

Okamura, T. and Sancar, A., Heelis, P.F., Begley, T.P., Hirata, Y., and Mataga, N.
J. Am. Chem. Soc. 1991, 113, 3143.

23

 

20.

21.

22.

23.

24.

25.

26.

27.

28.

29

30

31

32

Payne, B., Heelis, P.F., Rohrs, BR, and Sancar, S. Biochemistry 1987, 26, 7121.

Sancar, G.B., Jorns, M.S., Payne, G., Fluke, D.J., Rupert, CS, and Sancar, A.
J Biol. Chem. 1987, 262, 492-498.

Heelis, P.F., Okamura, T. and Sancar, A. Biochemistry 1990, 29, 5694-5698.
Li, Y.F., Heelis, P.F., and Sancar, A. Biochemistry 1991, 30, 6322-6329.

Kim, S.T., Sancar, A., Essenmacher, C., and Babcock, G.T. Proc. Natl. Acad. Sci.
USA 1993, 90, 8023-8027.

Hartman, RF. and Rose, S.D. J. Am. Chem. Soc. 1992, 114, 3559.

Ghisla, S., Massey, V., Lhoste, J ., and Mayhew, S.G., Biochemistry 1974, 13, 589-
597.

Payne, G. and Sancar, A., Biochemistry 1990, 29, 7719.

Hoffman, B.M. Roberts, .1 .E., Kang, CH, and Margoliash, E. J. Biol. Chem. 1987,
262, 1987.

. Essenmacher, C.E., Kim, S.T., Atamian, M., Babcock, GT, and Sancar, A. J. Am.
Chem. Soc. 1993, 115, 1602-1603.

. McCord, E.F., Bucks, RR, and Boxer, S.G., Biochemistry 1981, 20, 2880-2888.
. Heelis, P.F., Hartman, R.F., and Rose, S.D. Chem. Soc. Rev. 1995, 24, 289-297.

. Heelis, P.F., Hartman, R.F., and Rose, S.D. Chem. Soc. Rev. 1995, 24, 289-297.

Chapter 2: Isolation and Characterization of Photolyase

 

Exposure of DNA to ultraviolet radiation causes cyclobutane pyrimidine dimers
to form on the DNA strand. If left unrepaired, pyrimidine dimers can cause mutation,
cancer, and death.1 Photoreactivation is mediated by DNA photolyase which binds to
DNA-containing pyrimidine dimers independent of light and splits the dimers upon
exposure to near-UV light.2 To be able to study effectively the mechanism of action of
photolyase, large quantities of the enzyme are needed. Since very low concentrations of
photolyase are found in cells (10 to 20 molecules per cell in E. coli) the study of this
enzyme was difficult until 1978 when the photolyase gene was cloned3 and a photolyase
over-producing strain of E. coli was constructed in 1983.4 Sancar et al. were successful in
constructing a photolyase over-producing plasmid by joining a tac promoter to the
photolyase apoenzyme gene, phr. In this strain, photolyase constituted 15% of total
cellular proteins and the tac-phr plasmid amplified E. coli DNA photolyase
approximately 15,000-fold in the cell. By using this strain carrying the over-producing
plasmid, several milligrams of enzyme could be purified per gram of cells.

The buffer conditions (50mM Tris-HCI at pH 7.4) used in the lab of A. Sancar in
the purification procedure left the enzyme highly susceptible to air oxidation and
denaturation. The isolation procedure involved sonication, centrifugation, three dialysis
steps, an affinity column purification, gel filtration, and an hydroxyapatite column, all of
which lengthened the isolation procedure to several days.5 Owing to the time required for
this procedure, the enzyme had a greater chance of reacting with oxygen, causing the

oxidation state of the final product to be a mixture of semiquinone and oxidized states.

24

 

25

Since knowing the oxidation state of the enzyme is crucial to studying the mechanism of

repair, this isolation procedure was unsatisfactory and improvement was necessary.

MATERIALS AND METHODS

Bacterial Strains and Plasmids

 

Escherichia coli CSR603 was obtained from the E. coli Genetic Stock Center, Yale
University. M809 is identical to CSR603 with the exception that it is carrying the tac-
phr plasmid pMS969, the DNA photolyase strain. M809 was obtained from A. Sancar at
the North Carolina School of Medicine, Chapel Hill.

Chemicals

Isopropyl-B-D-thiogalactopyranoside (IPTG) used to induce the cells was obtained from
Boehringer Mannheim GmbH. The RNase and DNase used to cleave the RNA and DNA
strands were obtained from Sigma. The column materials used in the purification steps of
the isolation were Blue Sepharose CL-6B and Heparin Sepharose CL6B obtained from
Pharmacia LKB Biotechnology. The desalting columns packed with P-6DG
polyacrylamide gel (exclusion size of 6000 D) were obtained from Bio-Rad. The SDS-
PAGE molecular weight standard was from Bio-Rad.

m

The lysis buffer was made up of 50 mM Hepes, pH 7.0 with 100 mM NaCl, 10%(w/w)
sucrose, and 10 mM 2-mercaptoethanol. Buffer A consisted of 50 mM Hepes, pH 7.0,

10%(v/v) glycerol, and 10 mM 2-mercaptoethanol. Buffer B and Buffer C were identical

 

26
to Buffer A with the addition of 0.1 M KCl to Buffer B and the addition of 2.0 M KCl to

Buffer C.

Equipment

The French press used to break the cell walls in the initial step was obtained from SLM
Aminco, Urbana, IL. The optical spectra were obtained by using an AVIV
Spectrophotometer, Model 14DS UV-VIS, Lakewood, NJ. The centrifugations were
accomplished with the use of a Dupont Sorvall RC-SB Refrigerated Superspeed
Centrifuge, Hoffman Estates, IL. The SDS-polyacrylamide gels were produced by
utilizing a Bio-Rad Mini-PROTEAN II Electrophoresis Cell, Hercules, CA.

Assay of DNA Photolyase

 

The activity assay was performed as described in Jorns et al.6 by H. Sackett in the
Chemistry Department at Michigan State University. The substrate was poly dTlo DNA
obtained from the Biochemistry Department at Michigan State University and the host
was a concentrated stock solution of photolyase enzyme. The DNA was damaged and
repaired by using a UVP model UVGL-58 (V WR Scientific) combination
longwave/shortwave UV lamp. Photolyase was then added in the inactive semiquinone

form and was activated to the fully reduced form by being exposed to 366nm light.

RESULTS

Cell Growth

 

The cells were grown in Luria broth, which consists of 25g of Luria Broth powder (10

grams of tryptone, 10 grams of NaCl, and 5 grams of Yeast extract) per one liter of water.

 

 

27
The flasks were shaken at a speed of 250 rpm at 38°C. After the cells reached an optical

density of A600 = 0.65 to 0.8 (after about 6 to 8 hours), 2mM isopropylthio-b-D-
galactoside was added to induce the cells thereby causing the repressor on the DNA
strand to dissociate. This allowed the lac operon genes to be expressed and photolyase
concentration to be increased in the cell. The cells then continued to grow to reach log
phase (16 to 18 hours). The cells were harvested by centrifugation for 15 minutes at
4500rpm in a GSA rotor at 4°C. The cells were resuspended on ice in lysis buffer with
approximately 3ml of buffer per liter of culture. The suspension was homogenized and
stored at -80°C in a Sarstedt tube. Each liter of culture yielded about 3.5 grams of cells
(wet weight).

Isolation

The Sarstedt tube containing approximately 12 ml of suspended cells was thawed
in room temperature water, which took approximately 15 minutes. The cells were broken
by passing them through a French pressure cell two times at 20,000 psi and 4°C. To
remove any unbroken cells and cell fragments the lysed cells were centrifuged for 30
minutes at 19,000 rpm in an SS-34 rotor at 4°C. The clear brown supernatant containing
the photolyase was decanted from the pellet and its volume measured.

Three crystals each of DNase and RNase were added to the supernatant to cleave
the unwanted DNA and RNA strands further. This incubation was carried out for half an
hour with gentle stirring at 4°C. To precipitate the enzyme, ammonium sulfate (0.43 g per
ml of solution) was slowly added with stirring. After all of the ammonium sulfate had

dissolved, the suspension was centrifuged for 20 minutes at 19,000 rpm in an 88-34 rotor

 

28
at 4°C. The pellet was resuspended in approximately 30ml of buffer A. Desalting

columns obtained fiom Bio-Rad were used to remove the ammonium sulfate from the
protein solution. The desalting columns were equilibrated with buffer A and then 3ml of
protein solution was added to each of ten desalting columns. To wash the protein from
the colurrms, 4ml of buffer A was added to each column and the brown eluent was
collected.

A blue sepharose column (3 cm by 10 cm) was equilibrated with buffer A and the
eluent, diluted three-fold with buffer A, was loaded on to the column. The column was
rinsed with buffer A until two or three clear tubes had been collected. The enzyme was
then eluted with buffer C and the tubes with a blue color were combined. The protein
was then loaded onto desalting columns to remove the KC] in the eluting buffer. The salt
free solution was loaded onto a heparin column (2 cm by 10 cm) equilibrated with buffer
A. The column was rinsed with three column volumes of buffer A and the protein eluted
with a linear gradient composed of 100 ml of buffer B and 100 m1 of buffer C. The blue
fractions were combined and concentrated with the Amicon Ultrafiltration device with a
PM-10 membrane. Glycerol was added to give a final concentration of 30%(v/v) and the
protein was divided into lml aliquots and stored at -80°C in Eppendorf tubes. A flow
chart of this isolation procedure is shown in Figure 2.1. Photolyase is isolated with
greater than 95% purity as can be seen from the SDS-PAGE shown in Figure 2.2. An
absorbtion spectrum of the isolated enzyme can be seen in Figure 2.3 where absorbance at

580nm can be used to determine the final concentration of photolyase isolated

(8530:4800M"cm").

29

Figure 2.1. Flow chart of procedure for isolating photolyase.

 

30

Isolation Procedure

French press
centrifugation fior 30 minutes
digestion of DNA and RNA strands
enzyme precipitation ((NH4)ZSO4)
centrifugation for 20 minutes
pellet resuspension
desalting columns
Blue Sepharose column purification
desalting columns
Heparin Sepharose column purification

concentration

storage at -80°C

Figure 2.1

31

Figure 2.2. A Coomassie Blue-stained SDS-PAGE gel of photolyase.
Lanes 1 and 8, molecular mass standards (from top to bottom): ovalbumin
(45 kDa), trypsin anhydrase (21.5 kDa), aprotinin (6.5 kDa); lane 2,
unbroken E. Coli cells; lane 3, after French press; lane 4, after DNase and
RNase incubation; lane 5, after salt precipitation; lane 6, after blue
Sepharose column; lane 7, after white Sepharose column. Each lane
contained lSuL of protein.

32

 

 

 

 

 

Figure 2.2

33

Figure 2.3. Absorption spectrum of photolyase in its semiquinone form.

 

Absorbance

2.00

1.75

1.50

1.25

1.00 -

0.75 -

0.50

0.25

0.00

 

34

Wavelength (nm)

Figure 2.3

*10

35
Activity of the Purified Enzyme

 

The isolation procedure described in this chapter was made significantly more time
efficient by reducing the number of steps involved in the isolation relative to the number
of steps in the Sancar procedure. The buffer conditions were also altered in order to make
the enzyme more stable and therefore less susceptible to air oxidation. An activity assay
was conducted to ensure that the new buffer conditions did not impair the activity of the
enzyme. A stock solution of poly(dT) (2x10‘5M) in 50mM Hepes, pH 7.0, 100mM NaCl,
10mM 2-mercaptoethanol, and 10%(w/w) glycerol was irradiated in a stoppered quartz
semimicro cuvette (500 uL/cuvette). The cuvette was laid horizontally and irradiated at a
distance of 4cm from above by a short (1»=254nm) UV lamp for one minute intervals at
4°C. The extent of dimer formation was monitored by the decrease in absorbance at 260
and 290nm after each minute. When there was no longer any change in absorbance,
illumination was stopped.

The DNA stock solution was diluted to approximately 4.2x10'5M in thymine
concentration. The enzyme (1x10'7M) was added and the mixture was placed in a
stoppered semimicro cuvette with a final volume of lml. The cuvette was illuminated in
a horizontal position with the long (k=366nm) wavelength lamp (covered with a
watchglass to filter out radiation below 300nm) at 25°C. After each interval the
absorption at 260, 290, and 400nm was noted until the absorption readings remained
unchanged. The procedure was repeated with 50mM Tris-HCl at pH 7.2. As Figure 2.4

demonstrates, the activity of the enzyme was comparable in HEPES and Tris buffers.

36

Figure 2.4. Activity assay of photolyase in HEPES buffer and in Tris
buffer at 25°C.

M)

Concentration of Monomer (

 

Activity Assay Comparing
HEPES and Tris buffers

 

7.5x10'5 .
1 x tris
- HEPES
6 XXBT».
7.0x10— ”xx.
X9
_ XOX.
X0
6.5x10'5- >°
§
§
6.0x10'5— "
5
' t
i
5.5x10'5— an:
!
I
5.0x10'5..r,1,.,.l
o 10 20 30 40 50

Time (minutes)

Figure 2.4

1“.

38
DISCUSSION

By using the isolation procedure described here, photolyase enzyme could be
obtained with comparable purity to the procedure developed in the lab of A. Sancar. In
addition to obtaining a product that was 95% pure, this procedure was more time efficient
thereby allowing for a more accurate characterization of the enzyme and also giving a
greater product yield. The isolation procedure was made more time efficient by utilizing
techniques that accomplished the same task as in previous protocols, but in a shorter time
period. The number of steps required for purification of the enzyme was also reduced,
which decreased product loss and increased the yield by 92%. The breaking of the cells
was accomplished by utilizing a French press in place of sonication, which allowed a
greater percentage of cells to be broken effectively. Desalting columns were substituted
for dialysis, which resulted in less time needed to remove the salt from the eluting buffer.
Two affinity chromatography columns, which bound the photolyase while allowing the
impurities to pass through the column, were used in the final purification steps of the new
procedure. Photolyase was then eluted with a salt gradient. Because this procedure was
much quicker and was able to be performed at 4°C, the enzyme was much more stable in
that it was not as easily oxidized, which allowed us to isolate the protein in its blue
semiquinone form. The catalytic and spectroscopic properties of the enzyme isolated by

the procedure described here are comparable to literature values.

39

 

References

. Harm, W. Biological Eflects of Ultraviolet Radiation, Cambridge University Press,
London, 1980.

. Sancar, A. and Sancar, G.B. Annu. Rev. Biochem. 1988, 5 7, 29.

. Sancar, A. and Rupert, C.S. Gene 1978, 4, 294.

. Sancar, G.B., Smith, F .W., and Sancar, A. Nucleic Acids Res. 1983, 11, 667-678.

. Sancar, A., Smith, F.W., and Sancar, G.B. J. Biol. Chem. 1984, 259, 6028-
6032.

. Jorns, M.S., Sancar, GB, and Sancar, A. Biochemistry 1985, 24, 1856-1861.

Chapter 3: Identification of the transient, spin polarized EPR
signal of photolyase

 

 

When DNA photolyase is isolated in vitro, one electron oxidation of the fully
reduced active form to the inactive semiquinone form occurs. It has been proposed that
the flavin is reduced to its active form, FADH', by trp-306 that is located more than 13 A
away from the flavin.1 The oxidized tryptophan is then reduced by an exogenous
reducing agent. When there is no external reducing agent present, the flavin transfers the
electron back to the tryptophan within 10ms.2 A proposed mechanism for activation of
photolyase is shown in Figure 3.1.3 The fully reduced active form of the enzyme
catalyzes the repair of cyclobutadipyrimidines, which are created by exposure to
ultraviolet radiation (200 to 300nm), by using a light-induced electron transfer
mechanism.“‘5

A mechanism of dimer cleavage by photolyase has been suggested from the
crystal structure solved by Deisenhofer et al.6 From this crystal structure, it was proposed
that the pyrimidine dimer flips out of the DNA helix into a hole that leads to the binding
site found in the flat surface of photolyase. Once the dimer is bound to the enzyme, the
absorption of a photon initiates the repair mechanism. The folate molecule absorbs an
incident photon and subsequently the energy is transferred to the fully reduced flavin
molecule. The flavin, now in its excited state, donates an electron to the pyrimidine
dimer to generate a pyrimidine radical anion and FADH". The radical anion then splits
into monomers while transferring an electron back to the flavin.

Understanding the electron transfer processes described above is essential to

studying the mechanism of DNA repair. In order to study effectively these electron

40

41

Figure 3.1. Activation of photolyase by electron transfer from trp-306.

42

FADH} hV FADH?
-———————>-

<1———————
0 o +
11133 0 6 recomblnatlon 111330 6

exogenous
donor

FADH'

W%%

Heelis, P.F., Okamura, T. and Sancar, A. Biochemistry 1990, 29, 5694.

Figure 3.1

43

transfer processes, the oxidation states of the flavin must be known and controlled. Each
oxidation state has distinct absorption spectra as discussed in Chapter 1 and therefore can
be used to identify the oxidation state of the sample. In this study, the identity of the
species responsible for an observed transient EPR signal is investigated. This species
was previously reported as originating from the semiquinone form of the enzyme, but the
better understanding and control of the enzyme that we have obtained through the
biochemical approaches described in Chapter 2 has led to a different conclusion that is

discussed below.

Materials and Methods

Isolation of photolyase was performed as described in Chapter 2. Experiments
were carried out with protein samples that contained 50mM HEPES pH 7.0, 0.10M NaCl,
50% glycerol, 700uL/L BME and 3mM ferricyanide. The isolated enzyme in its
semiquinone form is relatively resistant to air oxidation and therefore requires the
addition of ferricyanide to oxidize the flavin cofactor. Prior to each experiment an
absorption spectrum was taken of the enzyme to verify the oxidation state by using an
Aviv 14DS UV-VIS Spectrophotometer (Lakewood, NJ). EPR measurements at 0°C
were performed with a TM011 cavity in a Varian E—4 that was interfaced to a personal
computer. The sample was held in the cavity by a TM quartz flat cell (Wilmad, Buena,
NJ). The frequency of the cavity was monitored with an EIP Microwave, Inc. (San Jose,

CA) frequency counter.

44

Photoexcitation of the sample was accomplished by using a Xenon flashlamp with
a l7ps pulse and a 50] lamp energy or a Quanta-Ray DCR-ll pulsed Nd-YAG laser
(Spectra-Physics, Mountain View, CA) equipped to provide 532 or 355nm light pulses
with a width of 10ns. Steady state EPR spectra were collected with a Metrabyte WAAG
data acquisition board to digitize the analog signal from the spectrometer. Kinetic traces
were collected by the use of computer software with data acquisition and timing
controlled by the Metrabyte WAAG board and a Metrabyte CTM-05 timing board.7 The
EPR instrument had a lOOKHz modulated detection system that provided an instrument
risetime of about 30ps . Transient EPR spectra were obtained by using an internally
triggered Stanford Research Systems SR 250 gated integrator and boxcar averager (Palo
Alto, CA).

Results

Figure 3.2 shows a steady state EPR spectrum of photolyase in its semiquinone
inactive form. The spectrum shows no well-resolved hyperfine couplings and a peak to
peak width of ~20G. It has an apparent g-value of 2.0039, which is consistent with
reports of flavin radicals that occur in proteins (20030—20040).8 Flash photolysis with
the Xenon flashlamp was conducted on the sample and produced an EPR transient
spectrum as shown in Figure 3.3. The transient spectrum was collected for 48us
following a delay of 4us after the lamp was triggered. The spectrum shows three major
peaks in a 1:2:1 intensity ratio each split by 15G. Superimposed on these 15G splittings
are smaller ones with 5G splitting. The kinetic trace that corresponds to the transient

spectrum is shown in Figure 3.4. The rise and fall of the light-induced radical occurs

45

Figure 3.2. Steady state spectrum of photolyase in its semiquinone form at
a concentration of 62.5uM. Experimental conditions were: 2.0 mW
microwave power, 9.22606 GHz microwave frequency, 30 1,18 time
constant, 3290 G center field, 100 G sweep width, 2.5 G modulation
amplitude at 25°C.

45

Figure 3.2. Steady state spectrum of photolyase in its semiquinone form at
a concentration of 62.5uM. Experimental conditions were: 2.0 mW
microwave power, 9.22606 GHz microwave frequency, 30 us time
constant, 3290 G center field, 100 G sweep width, 2.5 G modulation
amplitude at 25°C.

dX/dH

46

 

~80—

 

l

I l I 1 1 l r
3240 3260 3280 3300 3320

Field (6)

Figure 3.2

4
3340

47

Figure 3.3. Transient EPR spectrum of photolyase in its oxidized form at
a concentration of 73.5 pM. Experimental conditions were as follows: 6.3
mW microwave power, 9.39908 GHz microwave frequency, 2.5 G
modulation amplitude, 30 us time constant, and 100 G sweep. Boxcar
conditions were: 40 us delay, 435 us window, 15 one hour scans, and a
flash interval of 600 ms at a temperature of 4°C.

dX/dH

48

 

l 1 l r l l l

3320 3340 3360 3380
Magnetic Field (G)

Figure 3.3

 

49

Figure 3.4. Kinetic trace of the transient radical with photolyase in its
oxidized state at a concentration of 84.8uM and 200 flashes averaged.
Experimental conditions were: 6.3 mW microwave power, 9.411315 GHz
microwave frequency, 3350 G microwave field, 30 us time constant and
2.5 G modulation amplitude at a temperature of 25°C.

amplitude

 

 

l I I I

0 500 1000 1500

2000
thne(us)

Figure 3.4

 

117

51

with a half time of approximately 35113. This decay time, however, does not accurately
represent the lifetime of the radical due to limited instrument response time.

Previous reports stated that the steady state and transient signals arose from the
semiquinone form of the enzyme."l0 Figure 3.5 shows the mechanism for photoreduction
with the spin polarized species proposed to give the transient spectrum being
3‘FADH‘ --- trp'+.ll The enzyme samples used in those experiments were obtained from
A. Sancar and were a mixture of semiquinone and oxidized protein in ratios that varied
from sample to sample. Due to this mix of states, the assignment of the transient EPR
signal to the semiquinone form alone is difficult because either state, oxidized or
semiquinone, may contribute to this signal. The possible contribution from the oxidized
form of the enzyme was not considered in these experiments and therefore an incorrect
assignment was made. Evidence for this statement will be given in the following
paragraphs.

By using samples obtained via the isolation procedure described in Chapter 2, a
more accurate assignment of the species giving the EPR spectra could be made because
there was no longer a mixture of oxidation states. The samples used in the following
experiment were blue in color and made up of mainly the semiquinone form of the
enzyme, as determined by its absorption spectrum (see Figure 3.6). The semiquinone
form of photolyase has absorption maxima at 480, 580, and 625nm. The concentration of
the sample was determined by its absorbance at 580nm (a=5000M"cm") and found to be
13.5pM. By using this sample, an intense steady state EPR signal could be detected as

shown in Figure 3.7. This signal was identical to the signal reported by Essenmacher et

52

Figure 3.5 Proposed reaction mechanism for photoreduction that includes
the spin polarized species thought to give the transient EPR signal.

53

F ADH- —_ trp steady state

2 ( 3*FADH' — trp°+) <—> 4 ( 3*FADH' — trp-+)
spin polarized species
triplet quenching

FADH'

 

trp .+

l recombination

FADH-

 

trp

Essenmacher, C. PhD thesis 1995, Michigan State University.
Figure 3.5

 

 

54

Figure 3.6. Absorption spectrum of photolyase in its semiquinone form.
The concentration of the sample was 13.5 uM. The spectrum was obtained
by using an AVIV Spectrophotometer, Model 14DS UV-VIS at 25°C.

absorbance

0.25 —

0.20

0.15

0.10

0.05

 

0.00
300

Semiquinone Photolyase

 

I r I l l

400 500 600 700

wavelength (nm)

Figure 3.6

56

al.“'9 They also reported a transient signal and concluded that this signal originated from
the semiquinone state of the enzyme. We used the same sample that 'we had used for
the steady state experiment above and used similiar instrument parameters as
Essenmacher, et al. to try to reproduce their results, but we were not able to detect such a
transient signal with our sample.

A sample of yellow, oxidized protein was then used. The sample was oxidized
via the addition of 3mM ferricyanide to enzyme in the semiquinone form, which was then
air oxidized for twenty four hours. Before the addition of ferricyanide the ratio of
semiquinone to quinone was 1.37 and after oxidation was 0.157, indicating that most of
the sample was now in its oxidized state, as can be seen by its absorption spectrum shown
in Figure 3.7. The oxidized form of photolyase has an absorption maximum at 450nm
from which its concentration can be determined (8:1 1,300 M'lcm'l). Upon exposure to
flashes of light, the kinetic trace could be generated with this oxidized protein sample.
From this kinetic trace, a transient EPR signal was obtained identical to the transient
signal reported by Essenmacher et al. 8‘9, the only difference being that our sample was
fully oxidized while their sample contained both semiquinone and oxidized photolyase.
Considering the mixed oxidation state of the sample used in the previous study by
Essenmacher et al. , a conclusion can be made as to why the results differ from the results
presented here. The portion of sample in the semiquinone form gives the steady state
spectrum while the oxidized portion gives the transient signal. The transient signal is
only generated when light with wavelengths in the range of 400-450nm is used. No

signal could be observed when wavelengths above 500nm were used. Wavelengths

57
below 500 nm were filtered out by using a 500nm high pass filter with the Xenon

flashlamp. Since a transient signal is only observed when using wavelengths near the
absorption maximum of the oxidized protein and not the absorption maximum of the
semiquinone, the conclusion can be made that the transient signal arises from the
oxidized form of the enzyme.

Another experiment was conducted to observe the relationship between the
amplitude of the steady state and transient signals.‘2 If both signals originate from the
same species then their amplitudes should increase or decrease proportionally with the
change in semiquinone concentration. To perform this experiment, ferricyanide was
added to a 55 pM solution of photolyase to a final concentration of 3mM. A steady state
spectrum was taken and the amplitude of the signal noted. Then a kinetic trace was
obtained immediately after obtaining the steady state spectrum by giving the sample 500
flashes at 3349G and its amplitude noted. Every fifteen minutes this procedure was
repeated. As can be seen in Figure 3.8, the steady state signal decreases over time,
indicating that the sample is being oxidized, as the kinetic trace amplitude increases.
Therefore the two EPR signals, steady state and transient, cannot originate from the same
species. From these results and the experiments outlined above, it can be concluded that
the transient EPR spectrum originates from the oxidized, not the semiquinone form of
photolyase. A mechanism can be proposed, shown in Figure 3.9, with the fully oxidized
flavin as a member of the spin polarized pair. The oxidized flavin is excited and
intersystem crosses to the triplet state. The flavin triplet then abstracts an electron from

tryptophan. Acorrelated radical pair is formed consisting of a flavin doublet anda trp

58

Figure 3.7. Absorption spectrum of photolyase in its oxidized form. The
concentration of the sample was llSuM. The spectrum was obtained by
using an AVIV Spectrophotometer, Model 14DS UV-VIS at 25°C.

59

21)—

Oxidized Photolyase

absorbance

 

 

0_0 n . I . i . I
400 500 600 700

 

wavelength (nm)

Figure 3.7

 

60

Figure 3.8. Amplitude of transient signal versus amplitude of steady
state signal (see text for details).

 

Amplitude of Transient Signal

0.6 —

 

0.4

61

 

4O

1’

I ' I

60 80 150 140
Amplitude of Steady State Signal

1 1
100 160

Figure 3.8

62

Figure 3.9. Proposed reaction mechanism with the fully oxidized flavin as
a member of the spin polarized pair.

63

FAD— trp steady state

i hv
*FAD — trp

l intersystem crossing

*3FAD— trp

l electron transfer

2FAD‘— 2trp+

l recombination

FAD— trp

Figure 3.9

64

doublet producing the transient spin polarized signal. Recombination can then occur to

regenerate the original species.

65

 

References

1. Park, H., Kim, S.T., Sancar, A., and Deisenhofer, J. Science 1995, 268, 1866-1872.
2. Heelis, P.F., Payne, G. and Sancar, A. Biochemistry 1987, 26, 4634-4640.

3. Heelis, P.F., Okamura, T. and Sancar, A. Biochemistry 1990, 29, 5694.

4. Sancar, A. Advances in Electron Transfer Chemistry 1992, 2, 215-272.

5. Heelis, P.F., Hartman, R.F., and Rose, S.D. Chem. Soc. Rev. 1995, 24, 289-297.

6. Park, H.W., Kim, S.T., Sancar, A. and Deisenhofer, J. Science 1995, 268, 1866-1872.
7. Hoganson, CW. and Babcock, G.T. Biochemistry 1988, 27, 5848.

8. Bolton, J .R. Biological Application of Electron Spin Resonance 1972, ed. Swartz,
H.M. (John Wiley and Sons, Inc.) p. 24.

9. Essenmacher, C., Kim, S.T., Atamian, M., Babcock, GT, and Sancar, A. J. Amer.
Chem. Soc. 1993, 115, 1602-1603.

10. Kim, S.T., Sancar, A., Essenmacher, C., and Babcock, G.T. Proc. Natl. Acad. Sci.
USA 1993, 90, 8023-8027.

11. Essenmacher, C. Ph.D. dissertation 1995, Michigan State University, East Lansing,
Michigan.

12. Gindt, Y. unpublished results.

Chapter 4: Isotopic labeling studies performed on photolyase

 

When DNA photolyase is isolated, it is oxidized to the inactive FADH' form of
the enzyme, as discussed in Chapter 1. Reactivation to the fully reduced form occurs
when photolyase is irradiated with visible light and an electron is donated from a nearby
amino acid.1 In Kim et al.2, it was proposed that trp-306 is the immediate internal
photoreductant of FADH'.

Using EPR spectrometry, a transient spectrum can be observed upon illumination
of photolyase, as discussed in Chapter 3. The amplitude and sense of the kinetic signal
observed in a time-resolved experiment at fixed field changes with applied magnetic
field. The overall spectrum consists of three components in a 1:2:1 ratio in an
emission/absorption line shape. Each of the three components is split by approximately
15G and superimposed upon them are smaller splittings of 5G. The emissive and
absorptive transitions suggest that spin polarization occurs between the flavin
chromophore and the tryptophan radical.3 Preceding the spin polarization process,
flavin, in its fully oxidized state, is photoexcited and then intersystem crosses to the
triplet state. The flavin triplet then abstracts an electron from trp-306. This electron
transfer produces a flavin doublet and a trp doublet that act as a correlated radical pair to
produce the transient spin polarized signal.

To confirm that tryptophan is the transient amino acid radical involved in the
photoreduction of the flavin molecule, the indole ring of tryptophan was deuterated.2
When deuterium is substituted for hydrogen, the hyperfine couplings due to deuterium

are one-sixth those of hydrogen.4 Therefore, isotopic labeling can be used to determine

66

67

which species is responsible for the transient EPR spectrum that is seen with photolyase.
Hyperfine couplings will not be resolved in the spectra in which an isotopically labeled
sample is used because the couplings due to dueterium would be too small compared to
the line width of the EPR transitions of photolyase. When the tryptophan residue was
deuterated at positions 2, 4, 5, 6 and 7 (see figure 4.1), the hyperfine structure collapsed,
but the three major components were still observable.S These results suggested that the
smaller 5G couplings were due to coupling to or-protons on the tryptophan ring. The
conclusion has been made that the hyperfine coupling specifically originates from the
C(2) a-proton and the N(l) nitrogen on the indole ring of trp-306.2

The next question that arises is the origin of the major three line pattern of the
transient EPR spectrum. Molecular orbital (MO) calculations by the Hijckel—McLachlan
technique predict a three line pattern arising from coupling to the C(3) B-methylene
protons of a tryptophan cation radical.° To test this prediction and to rule out any
contribution from the flavin radical, specifically labeled tryptophan at the B-methylene
position can be used. If the three line pattern does indeed arise from coupling to these [3-
methylene protons, then the structure will collapse upon deuteration, as in the case of the
5G splittings. In theory, both radicals could contribute to the spin polarized signal, but
evidence shows that only the tryptophan contributes to the signal observed.4 This
observation is troublesome because a contribution from the flavin doublet should also be
seen in the spectrum. Experiments performed on enzyme samples that have deuterated

tryptophan at the B-methylene positions would help to investigate this issue.

68

Figure 4.1. Numbering scheme for the tryptophan molecule.

70

The goal of the work described in this chapter has been to learn to grow
photolyase on well-defined growth medium whereby the labeled tryptophan could be
incorporated into the enzyme, and ultimately to perform EPR experiments on these
samples. At present, photolyase has been successfully grown on the M-9 medium
constitution (see materials and methods for composition) while maintaining its EPR
characteristics, as can be shown by the work done with the control sample described
below. Future work will involve growing photolyase on the M-9 medium containing
labeled tryptophan at the B-methylene position. Using this labeled sample, EPR
experiments can then be performed to test if the large 1:2:1 splittings originate from these
B-methylene protons of the tryptophan residue.

Materials and Methods

Escherichia coli (strain PLK983/pMS969) was grown in M-9 medium containing
labeled tryptophan. M-9 medium consists of 200ml M-9 salts (64g sodium phosphate,
15g potassium phosphate, 2.5g sodium chloride, and 5g ammonium chloride diluted to
1L), 50g/L casein acids (Sigma, product number 65072-006), 0.1M calcium chloride, 1M
magnesium sulfate and 600mL Milli-Q water. This strain is unable to produce its own
tryptophan, which allows for labeled tryptophan to be incorporated into the enzyme. M-9
medium is a minimal medium, which means that it contains the minimal amount of
components needed for the cells to grow. In addition, the minimal medium differs from
Luria broth in that it does not contain yeast extracts, which would be a source of
unlabeled tryptophan. When the M-9 medium is used, the cell growth is retarded by a

factor of three owing to the decreased concentration of nutrients.

71

A control sample was produced by growing cells in M-9 medium that contained
unlabeled tryptophan. Isolation of the enzyme was performed as described in Chapter 2.
Experiments were carried out with protein samples that contained 50mM HEPES pH 7.0,
0.10M NaCl, 50% glycerol and 700pL 2-mercaptoethanol. The enzyme was isolated in
its fully oxidized form; therefore, no addition of ferricyanide was required. Prior to each
experiment, an absorption spectrum of the enzyme was recorded to verify the oxidation
state of the flavin by using an Aviv 14DS UV-VIS Spectrophotometer (Lakewood, NJ).

The deuterated sample was grown on M-9 medium containing labeled tryptophan
at the B—methylene position. The photolyase strain used in this study is unable to produce
its own tryptophan, so it relies upon the medium to supply the tryptophan that it needs to
survive. Since the tryptophan in the medium is labeled at specific positions, the
deuterated tryptophan is incorporated into the enzyme. Mass spectrometry can be utilized
to confirm the incorporation of the labeled tryptophan. The labeled enzyme was isolated
and prepared under the same conditions as the control enzyme described above.

EPR experiments were performed on a Bruker ESP 300E spectrometer by using a
TM011 cavity at 0°C. The sample was held in the cavity by a TM quartz flat cell
(Wilmad, Buena, NJ). The control and labeled samples were excited with a 17ps pulse
from a xenon flashlamp that was operated at 50J lamp energy. Kinetic traces were
collected by using computer software with data acquisition and timing controlled by a
Metrabyte WAAG board and a Metrabyte CTM-05 timing board.7 The EPR instrument
had a 100kHz modulation detection system that provided an instrument risetime of

approximately 30ps. An automation routine was written that would allow a transient

 

72
EPR spectrum to be obtained with the Bruker ESP 300E. This routine was designed so

that the magnetic field could be fixed while signal averaging. The automation routine
written and stored using ESP 300E internal software is as follows:

READP <filename>

EMF 3460.0

DEL

DACQ

WRITE <filename>_exp*0
TP inc 1

PG inc 1

EMF inc 20.0

DEL

DACQ

WRITE <filename>_exp_0
TP inc 1

PG inc 1

EMF inc -18.0

15 LOOP 3 20

OO\IO\Ul-i>-in\)>—t

r—dr—Il—‘v—Ar—t
AWN—‘00

Results

Experiments were performed on the control enzyme grown on minimal M-9
medium. This enzyme sample was used so that any changes in the EPR spectrum of the
labeled species could be attributed to the effects of labeling and not of the growth
medium. As can be seen in Figure 4.2, the absorption spectrum of the control enzyme is
similar to the fully oxidized enzyme from cells grown on rich medium. The peak that is
indicative of the fully oxidized enzyme is present, but is unexplainably red-shifted to
480nm. Peaks at 580nm and 624nm are also present, indicating that the sample is
slightly contaminated with flavin in its semiquinone state. Even though the absorption
peak of the oxidized flavin is slightly shifted, this absorption spectrum provides evidence

that the control enzyme is not affected significantly by growth on the M-9 medium.

73

Figure 4.2. Absorption spectrum of the enzyme grown on M-9 medium
with unlabeled tryptophan. The concentration of the sample was 133 pM.
The spectrum was obtained by using an AVIV Spectrophotometer, Model
14DS UV-VIS at 25°C.

 

absorbance

 

0.1

 

74

1 I

I 1 l

 

l l
400 500 600 700

wavelength (nm)

Figure 4.2

75

Figure 4.3. Kinetic trace of the transient radical obtained with the control
enzyme sample in its oxidized state at a concentration of 0.133mM and
300 flashes averaged. Experimental conditions were: 6.3mW microwave
power, 9.78 GHz microwave frequency, 3475 G microwave field, 30 us
time constant and 2.5 G modulation amplitude at a temperature of 0°C.

0.2

0.0

amplitude

T

 

 

 

 

 

 

 

76

I 1 I 1 I I I

 

500 1000 1500 2000
time (us)

Figure 4.3

 

77

Figure 4.4. Kinetic trace of the transient radical obtained with the control
enzyme sample in its oxidized state at a concentration of 0.133mM and
500 flashes averaged. Experimental conditions were: 6.3mW microwave
power, 9.78 GHz microwave frequency, 3485 G microwave field, 30 us
time constant and 2.5 G modulation amplitude at a temperature of 0°C.

 

78

06-

04—

02-

amplitude

 

'1
00%}

 

 

0.2 l . .
0 5a) 1111) 15(1)
linens)
Figure 4.4

79

Nonetheless, the absorption spectrum provides an indication that the chromophore
complement of enzyme from E. coli grown on minimal medium is not identical to wild-
type enzyme grown on Luria broth.

EPR experiments were performed with this control enzyme sample. Figures 4.3
and 4.4 show kinetic traces of the control enzyme. The transient signal changes sign
when the applied magnetic field setting is varied. These results are similar to the results
obtained with the fully oxidized wild type enzyme as described in Chapter 3. Since the
kinetic traces from the control and original samples behave identically, the conclusion
can be made that the growth medium does not affect the transient EPR experimental
results. Therefore any changes in the EPR spectra that are observed with the labeled
species can be attributed to isotope substitution effects and not to a change in growth
medium.

Future Work

In order to prove further the integrity of the control sample, a transient EPR
spectrum must be acquired. In order to perform such an experiment with the Bruker ESP
300E, the automation routine described above can be used. This automation routine
allows a transient EPR spectrum to be attained by utilizing controlled increments of the
magnetic field, while photoexciting the sample and averaging several scans. Once this
transient spectrum is obtained by using the control sample, it can be compared to the
transient spectrum of the wild type enzyme described in Chapter 3. Since the single field
kinetic traces of the wild type and control enzymes were nearly identical in behavior, the

transient spectrum is be expected to also be similar. The transient spectrum will allow us

80
to test conclusively the hypothesis that the growth medium does not affect the

spectroscopic characteristics of the enzyme samples.

After successful deuteration of the sample has taken place by following
procedures outlined above, the sample can be used for EPR experimentation. The
experiments will be performed on the Bruker ESP 300E spectrometer with the
automation program described to capture a transient EPR spectrum with the deuterated
sample. If the major couplings that are split by 15G in the natural abundance spectrum
can no longer be detected, the conclusion can then be made that deuteration of the B-
methylene protons has caused these 1:2:1 splittings to collapse, since the labeling of this
sample is the only variance from the control sample. These results can be used to test the
prediction that the three major peaks observed in the wild type, natural abundance EPR
spectrum are due to coupling to the B-methylene protons.

In summary, the growth and characterization of the control enzyme has been
conducted and the results analyzed. Now the same procedures must be carried out on the
deuterated photolyase sample to determine the origin of the hyperfine coupling

responsible for the major splittings of the transient EPR spectrum.

 

81

 

._‘

N

U.)

:5

\1

References

. Heelis, P.F., Okamura, T., and Sancar, A. Biochemistry 1990, 29, 5694-5698.

Kim, S.T., Sancar, A., Essenmacher, C., and Babcock, G.T. Proc. Natl. Acad. Sci.
USA 1993, 90, 8023-8027.

. Norris, J .R., Morris, A.L., Thurnauer, M.C., and Tang. J. J. Chem. Phys. 1990, 92,

423 9-4249.

Essenmacher, C., Kim, S.T., Atamian, M., Babcock, GT, and Sancar, A. J. Am.
Chem. Soc. 1993, 115, 1602-1603.

. Hoffman, B.M., Roberts, J.E., Kang, CH, and Margoliash, E. J. Biol. Chem. 1981,

256, 6556-6564.

. Essenmacher, C. Ph.D. dissertation 1995, Michigan State University, East Lansing,

Michigan.

. Hoganson, CW. and Babcock, G.T. Biochemistry 1988, 27, 5848.