Research Collection

Doctoral Thesis

Nucleosome remodeling activities act on UV-damagednucleosomes and facilitate DNA-repair

Author(s): Gaillard, Hélene

Publication Date: 2002

Permanent Link: https://doi.org/10.3929/ethz-a-004358664

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ETH Library

Diss. ETHN0 14692

Nucleosome Remodeling

Activities Act on UV-Damaged

Nucleosomes and Facilitate DNA-

Repair

A dissertation submitted to the

Swiss Federal Institute of Technology (ETH) Zurich

For the degree of

Doctor ofNatural Sciences

Presented by

Hélène Gaillard

Dipl. Biologie II, University of Basel

Born June 10th, 1975

Citizen of Bullet, VD

Accepted on the recommendation of

Prof. Dr. Fritz Thoma, examiner

Prof. Dr. Wolfram Hörz, co-examiner

Prof. Dr. Ulrich Suter, co-examiner

Zürich, 2002

Table of contents

Table of contents

Table of contents I

List of Figures IV

List of Tables VI

Abbreviations VII

Acknowledgments X

Summary 1

Résumé 3

1. Introduction 5

1.1. Chromatin Structure 6

1.1.1. The Nucleosome 6

1.1.2. Nucleosome Positioning 6

1.1.3. Nucleosome Dynamics 7

1.2. Chromatin Remodeling 8

1.2.1. Histone Modifications 8

1.2.2. ATP Dependent Chromatin Remodeling 9

1.3. UV Damage Formation 16

1.3.1. UV-Photoproducts 16

1.3.2. UV Damage Formation 18

1.4. Repair of UV Lesions 20

1.4.1. Nucleotide Excision Repair 20

1.4.2. Photoreactivation 20

1.4.3. CPD Glycosylases 23

1.5. Repair in Chromatin 24

1.5.1. NER and Chromatin 24

1.5.2. Photoreactivation in Chromatin 25

1.5.3. Repair and Chromatin Remodeling 26

1.6. Aim of Project 28

I

Table of contents

2. Results 29

2.1. The ATDED-long Nucleosome 29

2.1.1. Characterization ofthe ATDED-long Nucleosome 30

2.1.2. Photoreactivation Is Modulated in ATDED-long Nucleosomes 35

2.1.3. The Rotational Setting of ATDED-long Nucleosomes Does not Change

upon UV-Irradiation and Photoreactivation 40

2.1.4. Repair ofNucleosomal CPDs Remains Inhibited in the Presence of

Additional Photolyase 40

2.2. ySWI/SNF Remodeling of ATDED-long Nucleosomes 45

2.2.1. Remodeling by ySWI/SNF Alters the Structure of Nucleosomes 45

2.2.2. Enhanced CPD Repair Following ySWI/SNF Nucleosome Remodeling .48

2.3. yISW2 Remodeling of ATDED-long Nucleosomes 55

2.3.1. yISW2 Alters CPD Repair 55

2.3.2. Effect of yISW2 on ATDED-long Nucleosomes 64

2.4. The ATDED-short Nucleosome 73

2.4.1. Characterization ofthe ATDED-short Nucleosome 73

2.4.2. Photoreactivation Is Inhibited in ATDED-short Nucleosomes 76

3. Discussion 81

3.1. CPD Formation and Repair in ATDED Nucleosomes 81

3.1.1. UV-Damage Formation and Nucleosome Stability 82

3.1.2. Nucleosomes Inhibit Photoreactivation 82

3.1.3. Photolyase Does not Induce Octamer Movements on ATDED-longNucleosomes 84

3.1.4. Nucleosomal Inhibition of Photorepair Cannot Be Overcome by SequentialAddition of Photolyase 85

3.2. Remodeling by ySWI/SNF Alters the Structure of Nucleosomal DNA and

Facilitates CPD Accessibility 86

3.3. Nucleosome Mobilization by yISW2 Influences CPD Repair 88

3.4. Is Chromatin Remodeling Involved in DNA Repair in vivo? 90

3.5. ySWI/SNF and yISW2 Remodeling: Different Mechanisms? 92

4. Materials and Methods 94

4.1. ATDED Subcloning 94

4.1.1. P18ATDEDandpl8ATDED-c 94

4.1.2. pGEM-ATDED-short 94

II

Table of contents

4.2. Preparation of the DNA Fragments 95

4.2.1. Purification of the ATDED-long DNA Fragments 95

4.2.2. Purification of the ATDED-short DNA Fragment 96

4.2.3. End-labeling of the DNA Fragments 96

4.3. Preparation of Nucleosome Core Particles 97

4.4. Nucleosome Reconstitution 97

4.5. Remodeling with ySWI/SNF 98

4.5.1. Competition of ySWI/SNF 99

4.6. Remodeling with yISW2 99

4.7. DNasel Footprint 100

4.7.1. Maxam-Gilbert Sequencing 101

4.8. Restriction Enzyme Digestions 102

4.9. UV Irradiation 102

4.10. Photoreactivation 103

4.11. CPD Analysis 103

4.12. Quantification of Nucleoprotein and Sequencing Gels 104

4.13. Gel Electrophoresis 104

4.13.1.Low Melting Agarose Gels 104

4.13.2.Nucleoprotein Gels 105

4.13.3. Sequencing Gels 105

4.13.4.SDS-PAGE Gels 105

5. References 107

Curriculum Vitae 126

m

List of Figures

List of Figures

1. Introduction

Figure 1-1 : The two major photoproducts generated by UV light. .17

2. Results

Figure 2-1

Figure 2-2

Figure 2-3

Figure 2-4

Figure 2-5

Figure 2-6

Figure 2-7

Figure 2-8

Figure 2-9

Figure 2-10:

Figure 2-11:

Figure 2-12

Figure 2-13

Figure 2-14

Figure 2-15

Figure 2-16

Figure 2-17

Figure 2-18

Figure 2-19

Figure 2-20

Figure 2-21

Figure 2-22

Figure 2-23

Figure 2-24

Figure 2-25

Figure 2-26

Schematic drawing of the ATDED-long nucleosome 29

Native nucleoprotein gel electrophoresis 30

ATDED-long nucleosome has a define rotational setting 31

ATDED-long nucleosome has a define rotational setting 32

Restriction enzymes accessibility assay 33

UV footprint of ATDED-long nucleosomes 34

Native nucleoprotein gel analysis (containing 10% glycerol) 35

Modulation of CPD repair in ATDED-long nucleosomes 36

Modulation ofCPD repair in ATDED-long nucleosomes 37

Quantification of CPD repair in the top strand of ATDED-longnucleosomes 38

Quantification of CPD repair in the bottom strand of ATDED-longnucleosomes 39

DNasel footprint after UV irradiation and during photoreactivation..41

Native nucleoprotein gel analysis (containing 10% glycerol) 42

Photoreactivation of ATDED-long nucleosomes 43

Quantification of CPD repair in ATDED-long nucleosomes 44

Native nucleoprotein gel electrophoresis (containing 10% glycerol) .45

Altered nucleosome structure upon remodeling by ySWI/SNF 46

Native nucleoprotein gel of nucleosomes remodeled by ySWI/SNF

(containing 10% glycerol) 47

UV-footprint of nucleosomes remodeled by ySWI/SNF 49

Remodeling of ATDED-long nucleosomes by ySWI/SNF 50

Photoreactivation of nucleosomes remodeled by ySWI/SNF 52

Quantification of repair in remodeled nucleosomes 53

Quantification of repair in remodeled nucleosomes 54

Native nucleoprotein gel analysis 56

Photoreactivation of nucleosomes remodeled by yISW2 57

Photoreactivation of nucleosomes remodeled by yISW2 58

IV

List of Figures

Figure 2-27: Quantification of CPD repair by photolyase 60

Figure 2-28: Quantification of CPD repair by photolyase 61

Figure 2-29: Quantification of CPD repair by photolyase 62

Figure 2-30: Quantification of CPD repair by photolyase 63

Figure 2-31 : DNasel footprint of nucleosomes remodeled by yISW2 65

Figure 2-32: DNasel footprint ofnucleosomes remodeled by yISW2 66

Figure 2-33 : Native nucleoprotein gel analysis ofnucleosomes remodeled by yISW267

Figure 2-34: DNasel footprint of yISW2 remodeled nucleosomes 68

Figure 2-35: DNasel footprint ofyISW2 remodeled nucleosomes 69

Figure 2-36: Restriction enzyme analysis of nucleosomes remodeled by yISW2...70

Figure 2-37: Analysis ofDNA and nucleosomes remodeled by yISW2 before and

after UV irradiation 71

Figure 2-38: UV-footprint ofDNA and nucleosomes remodeled by yISW2 72

Figure 2-39: Schematic drawing of the ATDED-short nucleosome 73

Figure 2-40: Native nucleoprotein gel electrophoresis (containing 10% glycerol) .74

Figure 2-41 : ATDED-short nucleosomes have a defined rotational setting 75

Figure 2-42: UV footprint of the ATDED-short nucleosome 76

Figure 2-43: Photoreactivation of ATDED-short nucleosomes 78

Figure 2-44 : Quantification of photoreactivation in ATDED-short nucleosomes...79

4. Materials and Methods

Figure 4-1 : SDS-PAGE analysis of histone proteins. .97

v

List of Tables

List of Tables

1. Introduction

Table 1-1: The SWI/SNF group 13

Table 1-2: The ISWI group 14

4. Materials and Methods

Table 4-1: Oligos for ligation mediated PCR 95

Table 4-2: Sequence ofthe ATDED fragments 96

VI

Abbreviations

A adenine

À Angstrom, 10"10 metres

ADN acide désoxyribonucléique

ADP adenosine diphosphate

AMP adenosine monophosphate

ARS autonomously replicating sequence

ATP adenosine triphosphate

BER base excision repair

bps base pairs

BSA bovine serum albumin

C cytosine

°C degree Celsius

cm centimeter

CPD cis-syn cyclobutane pyrimidine dimer

CS Cockayne's syndrome

Da Dalton

DNA desoxyribonucleic acid

dNTP desoxynucleotide triphosphate

dTTP thymidine desoxyribonucleotide triphosphate

DTT dithiothreitol

E. coli Escherichia coli

EDTA ethylene diamine tetra acetate

EtBr ethidium bromide

FADH" 1,5-dihydroflavin adenine dinucleotide

Fig. figure

G guanine

GG-NER global-genome NER

h hour

HAT histone acetyltransferase

HDAC histone deacetylase

8-HDF 8-hydroxy-5-deazariboflavin

HEPES hydroxyethylpiperazine ethanesulfonic acid

H20 water

VII

J

k

KCl

KMn04

L

M

m

mA

MDa

MgCl2

MMTV3'LTR

MTHF

MU

MWCO

H

n

NaAc

NaCl

NaOH

NEB

NER

Nucl.

P

pb

PCR

pH

Phr.

PMSF

(6-4)PP

RNA

rpm

rRNA

S. cerevisiae

SDS

SV40

joule

kilo, 103

potassium chloride

potassium permanganate

litre

molar

milli, 10"3

milli Ampère

mega Dalton

magnesium chloride

mouse mammary tumor 3 '

long terminal repeat

5,10-methenyltetrahydrofolate

map units

molecular weight cut off

micro, 10"6

nano, 10"9

sodium acetate

sodium chloride

sodium hydroxide

New England Biolabs

nucleotide excision repair

nucleosomes

pico, 10"12

paire de bases

polymerase chain reaction

potentia hydrogenii, concentration of [H30+]

photolyase

phenylmethylsulfonyl fluoride

pyrimidine (6-4) pyrimidone photoproduct

ribonucleic acid

revolutions per minute

ribosomal RNA

Saccharomyces cerevisiae

sodium dodecyl sulfate

simian virus 40

VIII

Abbreviations

T thymine

TBE 90 mM Tris-borate, 2 mM EDTA

TCA trichloroacetic acid

TC-NER transcription-coupled NER

TE 10 mM TRIS, 1 mM EDTA

T4-endoV T4 endonuclease V

tRNA transfer ribonucleic acid

TTD trichothiodystrophy

U unit

UV ultraviolet

V volt

W watt

w/v weight pro volume

XP xeroderma pigmentosum

IX

Acknowledgments

Acknowledgments

I would like to thank Fritz Thoma for teaching me a lot about science and for revealing me some

of my limits and qualities.

I thank Ulrich Suter for his support and for coexamining my thesis, and Wolfram Hörz for the

coexamination ofmy thesis.

Many thanks to: Craig Peterson for providing the ySWI/SNF complex and for his enthusiasm.

Corey Smith for purifying the ySWI/SNF complex and dispensing technical advice. And Tim

Richmond for providing the yISW2 complex.

Very special thanks go to Dan Fitzgerald for purifying the yISW2 complex, for his ideas, his

enthusiasm, his laugh, and the very good job he did on our manuscript.

A huge thank to Ralfi Wellinger for valuable ideas and discussions, and for introducing me in the

football team.

'Gracias' to Andres Aguilera for lending me his office and to the departamento de genética for

providing me with a quiet space in the library, internet connection and a stimulating' latin'

atmosphere.

I acknowledge the Roche Research Foundation for allowing me a fellowship to finish my thesis,

and Cuno Künzler, Carla Zingg and Ursula Scheier for advises and encouragements.

Thanks to Andreas Meier, for his help and discussions, and to all the former and present

members ofthe Thoma and Sogo groups, especially Bernie Suter, Magda Livingstone, Christoph

Capiaghi, Teresa Lettieri, Rachel Jossen, Sandra Ursprung, Martin Burkhalter, Massimo Lopes

and José Sogo for their scientific input, nice chats and good atmosphere.

Many thanks to Bernd Walzel, Dietbert Neumann and David Genoux, for good scientific

discussions and for making work more enjoyable, and to all the members of the IZB for

providing me with an excellent place to work in.

Un grand merci à mes parents, grand-parents, à Françoise et à André, ainsi qu'au reste de ma

famille et à mes amis, pour ce qu'ils sont et pour leur soutient.

X

Summary

Summary

UV light induces damages in DNA, leading to structural distortions that impede basic

cellular functions such as transcription and replication. DNA lesions have to be repaired to

prevent mutations, cancer and cell death. Cyclobutane pyrimidine dimers (CPDs) are the major

DNA lesions generated by UV light. CPDs are repaired either by nucleotide excision repair

(NER) or by photoreactivation. NER is a multistep pathway conserved from yeast to human. For

photoreactivation, a single enzyme called photolyase uses light as energy source to repair CPDs.

The mechanism of photoreversion involves flipping-out the CPD into the active site of the

photolyase enzyme. In eukaryotic cells, DNA is folded in nucleosomes and higher order

chromatin structures. Nucleosomes exert a repressive influence on the biological functions of

DNA by restricting its accessibility to proteins. Despite of packaging in chromatin, CPDs are

completely repaired by NER and by photoreactivation in vivo. However, preceding work

demonstrated that a nucleosome positioned on a short DNA sequence (134 bps) severely inhibits

photoreactivation in vitro. Therefore, it is not known whether in vivo dynamic properties of

nucleosomes are sufficient for repair of nucleosomal CPDs or whether nucleosome remodeling

machines might be required. Here, nucleosome reconstitution and photolyase were used to

investigate how structural and dynamic properties of nucleosomes and chromatin remodeling

complexes contribute to damage recognition and processing.

We reconstituted a positioned nucleosome at one end of a 226 bps long DNA sequence

('ATDED-long') to investigate whether providing space for octamer sliding facilitates

nucleosomal photorepair. DNA-damage was induced by UV-light and repair was analysed by

photolyase. We observed slow and inefficient repair in nucleosomal DNA, whereas DNA outside

of the nucleosome was efficiently repaired. Despite of the length of the fragment, the

nucleosome was neither displaced by UV damage nor during photoreactivation. Thus, the

ATDED-long nucleosome mimics the in vivo situation, where repair by photolyase is modulated

by positioned nucleosomes. In contrast to the almost complete repair of nucleosomal lesions in

living cells, repair remains inhibited in the ATDED-long nucleosomes even after long incubation

times, suggesting that nucleosome disruption or displacement might be required for efficient

repair in vivo.

ATP-dependent chromatin remodeling complexes are molecular machines that use the

energy ofATP hydrolysis to alter the structure of nucleosomes and promote octamer sliding. The

ySWI/SNF and yISW2 remodeling complexes have been shown to play important roles in the

1

Summary

regulation of transcription. However, it is not known whether they also assist other DNA-

dependent processes, like replication, recombination and DNA repair. In this work we found that

both, ySWI/SNF and yISW2 were capable to remodel UV-damaged ATDED-long nucleosomes

and facilitated repair of nucleosomal CPDs by photolyase. While ySWI/SNF altered the

conformation of nucleosomal DNA and promoted more homogeneous repair, yISW2 altered the

repair pattern by moving the nucleosome to a more central position on the DNA fragment. Thus,

two different nucleosome remodeling complexes can act on UV-damaged nucleosomes and

facilitate repair by a 'flip-out' enzyme. It is therefore possible that similar activities are engaged

in living cells to relieve the inhibitory effect of nucleosomes for photoreactivation and other

DNA-repair processes.

2

Résumé

Résumé

Les rayonnements ultraviolets (UV) induisent des lésions provoquant des distortions

structurelles dans l'ADN qui interfèrent avec les fonctions cellulaires de base telles que la

transcription et la replication. Les lésions dans l'ADN doivent être réparées afin d'éviter

mutations, cancer et mort cellulaire. Les dimères cyclobutyliques de pyrimidines (CPDs) sont les

principales lésions engendrées par les UV. Les CPDs sont réparés par excision de nucleotides

(NER) ou par photoréactivation. La réaction de NER est un système complexe conservé de la

levure à l'être humain et impliquant plusieurs étapes. Pour la photoréactivation, une seule

enzyme appelée photolyase utilise la lumière comme source d'énergie pour réparer les CPDs. La

photolyase retourne la lésion dans son site actif pour la photoréversion (mécanisme de 'flip-

out'). Dans les cellules eucaryotes, l'ADN est assemblé en nucleosomes et en structures

chromatidiennes supérieures. Les nucleosomes exercent une action répressive sur les fonctions

biologiques de l'ADN parce qu'ils en limitent l'accès aux protéines. Malgré l'organisation de

l'ADN en chromatine, les CPDs sont complètement réparés par NER et par photoréactivation in

vivo. Cependant, des études antérieures ont démontré qu'un nucleosome positionné sur un

fragment d'ADN court (134 pb) inhibe fortement la photoréactivation in vitro. On ne sait pas si

les propriétés dynamiques inhérentes aux nucleosomes suffisent pour réparer les CPDs

nucléosomaux in vivo, ou si des machineries de remodelage de nucleosomes sont requises. Nous

avons utilisé la reconstitution de nucleosomes et la photoyase pour élucider comment les

propriétés structurelles et dynamiques des nucleosomes d'une part, et les complexes de

remodelage de nucleosome d'autre part, contribuent à la reconnaissance et à la réparation des

lésions.

Nous avons reconstitué un nucleosome positionné à une extrémité d'un fragment d'ADN

de 226 pb ('ATDED-long') pour tester si le fait de donner suffisament de place à l'octamère pour

'glisser' le long du fragment d'ADN facilite la photoréparation nucléosomale. Les nucleosomes

ont été exposés aux UV pour introduire des lésions dans l'ADN et leur réparation analysée avec

la photolyase. Nous avons observé une réparation lente et inefficace dans l'ADN nucleosomal,

alors que l'ADN situé en dehors du nucleosome était réparé efficacement. Malgré la longueur du

fragment, le nucleosome n'a été déplacé ni par les lésions UV ni pendant la photoréactivation.

Ainsi, le nucleosome ATDED-long mime la situation in vivo, où la réparation par

photoréactivation est modulée par des nucleosomes positionnés. Toutefois, contrairement à la

réparation pratiquement complète observée dans les cellules vivantes, la réparation reste inhibée

3

Résumé

dans le nucleosome ATDED-long même après de longues incubations, suggérant que la

disruption ou le déplacement du nucleosome pourraient être requis pour une réparation efficace

in vivo.

Les complexes de remodelage de chromatine ATP-dépendants sont des machineries

moléculaires qui utilisent l'énergie d'hydrolyse de l'ATP pour changer la structure des

nucleosomes et promouvoir le déplacement des octamères. Les complexes de remodelage ySWI/

SNF et yISW2 jouent des rôles importants dans la régulation de la transcription. Par contre, on ne

sait pas si ils assistent également d'autres processus impliqués dans le métabolisme de l'ADN,

comme la replication, la recombinaison et la réparation. Dans cette étude, nous avons montré que

ySWI/SNF et yISW2 sont tous deux capables de remodeler les nucleosomes ATDED-long

endommagés par les UV et de faciliter la réparation de CPDs nucléosomaux par la photolyase.

Alors que ySWI/SNF modifie la conformation de l'ADN nucleosomal et promouvoit une

réparation plus homogène, yISW2 change le schéma de réparation en déplaçant le nucleosome à

une position plus centrale sur le fragment d'ADN. Donc, deux complexes de remodelage de

nucleosome différents peuvent agir sur des nucleosomes endommagés et faciliter la réparation

par une enzyme utilisant un mécanisme de 'flip-out'. En conséquence, il est possible que des

machineries similaires soient engagées dans les cellules vivantes pour soulager l'effet inhibiteur

des nucleosomes pour la photoréactivation et d'autres systèmes de réparation de l'ADN.

4

1 Introduction

1. Introduction

Each cell is constantly challenged by a variety of DNA damaging factors. First,

environmental agents such as the ultraviolet (UV) component of sunlight, ionizing radiation and

numerous genotoxic chemicals cause alterations in DNA. Second, by-products of normal cell

metabolism constitute a permanent enemy to DNA from within. These include oxygen species

derived from oxidative respiration and products of lipid peroxidation (Davies, 2000). Finally,

some chemical bonds in DNA tend to spontaneously disintegrate under physiological conditions.

Hydrolysis of nucleotide residue leaves non-instructive abasic sites (Lindahl, 1993). If left

unrepaired, DNA lesions may lead to mutations, cancer or cell death. Many lesions block

transcription. Lesions also interfere with DNA replication. A growing class of specialized

translesions polymerases able to bypass DNA-lesions have been found, but translesion DNA

synthesis often comes at the expense of a higher error rate, subsequent mutations and cancer

(Bebenek and Kunkel, 2002; Friedberg et al., 2001). Cells arrest at various checkpoints in

response to DNA damage, allowing time for repair (Zhou and Elledge, 2000). Lesion detection

may occur by blocked transcription, replication or specialized sensors. When damage is too high,

a cell may opt for the ultimate mode of rescue by initiating apoptosis at the expense of a whole

cell (Rich et al., 2000). All cells have a variety of repair pathways, which are specialized in repair

of defined damages (Hoeijmakers, 2001b).

In eukaryotic cells from yeast to mammals, DNA is folded into the compact structure of

chromatin. Packaging of DNA into nucleosomes and chromatin fibers changes the structure of

the DNA and severely restricts its exposedness to proteins (Luger et al., 1997; Thoma, 1999;

Wolffe, 2000), affecting thereby DNA dependent processes. The nucleosome, in its role as the

principal packaging element of DNA within the nucleus, is the primary determinant of DNA

accessibility. Therefore, positioned nucleosome can act as regulators of transcription and

replication (Han and Grunstein, 1988; Knezetic and Luse, 1986; Lorch et al., 1987).

Molecular machines which affect packaging of DNA have been shown to play crucial

roles in the regulation of transcription (Narlikar et al., 2002; Wolffe and Guschin, 2000). In

analogy to transcriptional regulation, it has been proposed that such complexes might increase

the accessibility of DNA lesions, thereby facilitating repair (Fyodorov and Kadonaga, 2001;

Green and Almouzni, 2002; Meijer and Smerdon, 1999).

5

1. Introduction

1.1. Chromatin Structure

1.1.1. The Nucleosome

DNA in chromatin is organized in arrays of nucleosomes (Romberg, 1977). The DNA that

connects two adjacent nucleosome is referred to as linker DNA. The X-ray crystal structure of

the nucleosome core particle at 2.8 Â resolution (Luger et al., 1997) provided detailed insight in

the arrangement of the histones and the DNA and the path of the DNA helix. The nucleosome

core consists of 147 bps of DNA wrapped in 1.65 left-handed superhelical turns around an

octamer of core histones. The DNA binds the octamer with a central base pair at the pseudo-dyad

axis, which divides the nucleosomal DNA into 73 and 72 bps halves, with one base pair falling

on the dyad. The additional base pair is accommodated without disruption of histone-DNA

contacts (Luger et al., 1997). The whole nucleosome particle has a diameter of 11 nm in length

and 5.5 nm in width. The path of the DNA superhelix is not uniform but distorted owing to the

local structure of the histone DNA-binding surface. The overall helical periodicity of the DNA

around the nucleosome core is 10.2 bps compared to 10.6 bps in B-DNA (Hayes et al., 1990). A

set of contacts is made every 10 bps where the minor groove of the double helix faces inward.

Electrostatic interactions with the DNA phosphates, as well as nonpolar contacts with the

deoxyribose groups are observed.

The disc-shaped histone octamer contains two copies of each four histones H2A, H2B, H3

and H4 in a tripartite assembly consisting of a central (H3)2(H4)2 tetramer flanked by two (H2A-

H2B) dimers (Arents et al., 1991; Eickbush and Moudrianakis, 1978). The four nucleosome core

histones contain a conserved histone fold motif within their central DNA binding domain (Arents

and Moudrianakis, 1995). Each of the core histones has a highly positively charged N-terminal

region; and histones H2A and H3 have analogous domains at their C-termini as well. The N-

terminal tails of both H2B and H3 pass through minor groove channels, so there is a protruding

tail every 20 bps (Luger et al., 1997). H2A and H4 tails pass across the superhelix on the flat

faces of the particle to the outside as well. These 'tail' domains are known to be extended and

mobile, and are the site of numerous posttranslational modifications (see Section 1.2.1.;

(Jenuwein and Allis, 2001)).

1.1.2. Nucleosome Positioning

The position of a nucleosome on the DNA sequence is determined by its 'translationaP

and 'rotational' setting (Simpson, 1991; Thoma, 1992). Translational positioning describes the

location of the octamer on the DNA sequence, rotational positioning the orientation of the double

helix with regard to the octamer surface.

6

1. Introduction

In vitro, positoning of nucleosomes along the DNA molecule is mainly governed by DNA

sequence preferences (Wolffe, 2000). It is energetically favourable to incorporate a DNA

sequence that is already curved into a nucleosome, as DNA has to be bent around the histone

core anyway (Drew and Travers, 1985b; Shrader and Crothers, 1989). On the other hand,

extensive homopolymeric stretches of rigid oligo(dA)(dT) or hairpin DNA structures are not

favoured for incorporation into nucleosomes (Garner and Felsenfeld, 1987; Nobile et al., 1986;

Prunell, 1982). Thus, the local influences of DNA rigidity and curvature will influence the

position of the histone octamer along the DNA backbone. Once formed, a nucleosome may not

be entirely fixed in position (Flaus et al., 1996; Meersseman et al., 1992). The statistical

preference observed for the DNA minor groove to face the histone octamer at (A+T)-rich

sequences indicates that certain sequences will impart a rotational orientation to the DNA double

helix when bound in a nucleosome (Travers and Klug, 1987).

In vivo, parameters that affect the nucleosome position are sequence dependent bentability

of the DNA, proteins that position nucleosomes, and chromatin folding (Thoma, 1992; Widom,

1998).

1.1.3. Nucleosome Dynamics

Nucleosomes are dynamic structures undergoing structural transitions that affect the

accessibility of the DNA. The histone octamers may slide along the DNA (nucleosome mobility;

(Meersseman et al., 1992)). Multiple nucleosome positioning with unique rotational setting was

shown in vitro and in vivo for the sea urchin and S. cerevisiae 5 S rRNA genes, indicating that the

rotational information of DNA is a determinant of nucleosome positioning (Buttinelli et al.,

1993; Buttinelli et al., 1995; Di Marcotullio et al., 1998; Pennings et al., 1991). Nucleosomes

reconstituted on L. variegatus 5 S rRNA and MMTV 3'LTR sequences were shown to migrate

along DNA at low ionic strength within a conserved rotational phase (Flaus et al., 1996; Flaus

and Richmond, 1998). Generally, there seems to be a preference to maintain the rotational

setting, while translational setting is less strongly enforced. Nevertheless, nucleosomes may

wobble within a few base pairs, transiently exposing different nucleotides to the nucleosomal

surface (Tanaka et al., 1996; Thoma, 1991).

Nucleosome could also undergo natural dissociation/reassociation processes leading to

transient exposure of the DNA. Dissociation involves presumably release of two (H2A-H2B)

dimers prior to the dissociation of the (H3)2(H4)2 tetramer (Khrapunov et al., 1997; Yager et al.,

1989). Alternatively, nucleosomes may unfold and refold without dissociation of the histone

octamer. The structural transition can be generated in vitro at very low ionic strength (Thoma et

7

1. Introduction

al., 1979). Because of the multiple DNA-binding sites of the octamer, dissociation ofDNA from

the nucleosome would occur in semi-cooperative stages, beginning at the entry and exit points of

the DNA, and then proceed into the H2A-H2B dimer, and eventually into the H3-H4 tetramer

regions (Luger et al., 1997). For example, it has been shown that melting of DNA in the

nucleosome core particle is biphasic, and that the initial denaturation phase is due to melting of

the DNA termini (McGhee and Felsenfeld, 1980; McGhee et al., 1980; Seligy and Poon, 1978).

Both yeast RNA polymerase and exonuclease III were shown to proceed into the nucleosome

core by steps of about 10 bp (Prunell, 1983; Riley and Weintraub, 1978; Studitsky et al., 1997),

consistent with a sequential disruption of the histone-DNA contacts. Measurement of the

accessibility of restriction enzyme sites in nucleosome cores showed an increasing resistance to

digestion of the order of 102 for sites at DNA termini, to 105 for sites at the DNA centre,

compared to free DNA (Polach and Widom, 1995). At the molecular level, nucleosomal DNA is

thought to transiently unwrap and rebind to the histone octamer.

1.2. Chromatin Remodeling

Many recent studies have concentrated on identifying protein complexes capable of

unfolding or modifying chromatin structure. The most widely characterized chromatin-

modifying complexes studied to date can be classified into two major groups, based on their

mode of action. (1) Histone modifying complexes, which regulate the transcriptional activity of

genes by covalent modification of specific residues of the core histones (reviewed in (Grant,

2001; Green, 2001; Marmorstein, 2001; Schüssel, 2000)). (2) ATP-dependent complexes, which

use the energy ofATP hydrolysis to locally disrupt or alter the association of histones with DNA

(reviewed in (Narlikar et al., 2002; Peterson, 2000; Varga-Weisz, 2001; Vignali et al., 2000)).

1.2.1. Histone Modifications

Histones are the predominant protein components of chromatin. Each histone core protein

contains flexible and highly basic 'tails domains', which are highly conserved across various

species. Post-transcriptional modification of conserved amino acids, notably methylation, ADP-

ribosylation, phosphorylation, ubiquitination and acetylation, modulate the interaction potential

of the tail domains, and may influence the folding and functional state of the chromatin fiber

(Berger, 2001; Grunstein, 1997; Jenuwein and Allis, 2001; Marmorstein, 2001). Such histone

modification have long been correlated with various nuclear activities, including replication,

chromatin assembly and transcription (Durrin et al., 1991; Thompson et al., 1994). Histone

modifications may also play a role in signalling other chromatin-regulatory and transcriptional

8

1. Introduction

events. It has been proposed that distinct histone modification pattern might act sequentially or in

combination to form a 'histone code' that is read by other proteins to elicit distinct downstream

transcriptional events (Jenuwein and Allis, 2001; Strahl and Allis, 2000).

Histone acetylation is the most studied post-transcriptional modification and has formed

the basis for an evolving model for how histone modifications may modulate chromatin structure

(reviewed in (Eberharter and Becker, 2002)). Acetylation of internal lysine residues of core

histone N-terminal domains has been found correlatively associated with transcriptional

activation in eukaryotes for more than three decades (Allfrey et al., 1964; Gorovsky et al., 1973;

Mathis et al., 1978). However, the connection between histone acetylation and transcription

remained uncertain until it was demonstrated that yeast Gcn5 protein, a positive transcriptional

regulator ofmany genes, has histone acetyltransferase (HAT) activity (Brownell et al., 1996) and

stimulation of transcription by Gcn5 requires the HAT activity (Kuo et al., 1998). The

implication were soon reinforced by similar results on mammalian transcription factors

(reviewed in (Grunstein, 1997; Struhl, 1998; Wolffe and Guschin, 2000)). Almost in parallel with

the discovery of HATs, came the identification of histone deacetylase (HDAC) enzymes, whose

activities have been correlated with transcriptional repression (Pazin and Kadonaga, 1997;

Taunton et al., 1996; Wolffe, 1997). Histones are locally modified on target promoters and

specific lysines in particular histones are target for HATs and HDACs (Kuo et al, 1996; Zhang et

al., 1998a). Acetylation is also a global process, since entire chromatin domains are targeted for

acetylation (Hebbes et al., 1994). However, the mechanisms by which whole domains are

targeted for acetylation is currently not known.

1.2.2. ATP Dependent Chromatin Remodeling

Large protein complexes that use the energy of ATP hydrolysis to disrupt or alter the

association of histones with DNA have been purified from yeast, fly, frog, and human (reviewed

in (Aalfs and Kingston, 2000; Fyodorov and Kadonaga, 2001; Havas et al., 2001; Narlikar et al.,

2002; Peterson, 2000; Vignali et al., 2000). ATP-dependent complexes can move positioned

nucleosomes, thereby exposing or occluding DNA sequences, and can create conformations

where DNA is accessible on the surface of the histone octamer.

All of the ATP-dependent chromatin-remodeling complexes contain an ATPase subunit

that belongs to the SNF2 superfamily of proteins. Based on the identity ofthis subunit, they have

been divided into three groups, the SNF2, the ISWI and the Mi-2 groups (Boyer et al., 2000a).

Due to distinct biochemical properties of individual remodeling enzymes, it has been proposed

9

1. Introduction

that the enzymes of the SWI/SNF and the ISWI groups use distinct mechanisms to remodel

chromatin structure. However, the mechanism by which nucleosome remodeling is achieved

remains poorly understood.

Although functional analysis of these complexes has been restricted mainly to

transcription, it is likely that similar activities may also assist replication, recombination and

repair (Fyodorov and Kadonaga, 2001; Green and Almouzni, 2002; Narlikar et al., 2002).

1.2.2.1. The SWI/SNF Group

This group includes yeast SWI/SNF, yeast RSC, yINO80, the drosophila Brahma (BRM)

complex and human SWI/SNFs (BRG1 and hBRM). All of these contain an ATPase subunit that

is highly related to yeast SWI2. They hydrolyze ATP in the presence of DNA or nucleosomes

(Cairns et al., 1996; Cote et al., 1994; Kwon et al., 1994) and use the energy generated to perturb

nucleosome structure. Some features of the remodeled nucleosomes in vitro are described in

Tab. 1-1.

ySWI/SNF is 2 MDa in size and contains 12 subunits (SWI1, SWI2/SNF2, SWI3, SNF5,

SNF6, SNF11, SNF 12, SWP29, SWP73, SWP82, Arp9, and Arp7). In vitro studies have shown

that the ySWI/SNF complex disrupts the rotational phasing ofDNA on the surface of the histone

octamer (Cote et al., 1994; Owen-Hughes et al., 1996). Another consequence of ySWI/SNF

nucleosome disruption is enhanced accessibility of DNA-binding proteins to nucleosomal DNA.

This has been shown with several types of sequence specific DNA-binding proteins and with

restriction endonucleases (Cote et al., 1994; Logie and Peterson, 1997; Owen-Hughes et al.,

1996; Utley et al., 1997). In addition, ySWI/SNF has been shown to promote octamer sliding

along the DNA (Jaskelioff et al., 2000; Whitehouse et al., 1999). Neither the eviction or gross

rearrangement of H2A/H2B dimers nor the splitting of H3/H4 tetramer are required for

remodeling, since ySWI/SNF can remodel arrays that harbour tetramers and disulfide-linked

tetramers (Bazett-Jones et al., 1999; Boyer et al., 2000b).

In vivo, the ySWI/SNF complex has been shown to activate transcription of specific target

genes. It interacts directly with the activation domains of sequence specific acidic activators, like

Gcn4p, VP16 and Hap4, which recruits the complex to the target promoters (Natarajan et al.,

1999; Neely et al., 2002; Neely et al., 1999; Yudkovsky et al., 1999). A close collaboration

between the histone acetyltransferase complex Gcn5 and the remodeling factor ySWI/SNF in the

transcriptional activation of some genes has been shown (Biggar and Crabtree, 1999; Gregory et

al., 1999; Pollard and Peterson, 1997).

10

1 Introduction

Only a small fraction of yeast genes require ySWI/SNF for activation, and ySWI/SNF is

involved in repression of almost the same amount of genes (Holstege et al., 1998; Sudarsanam et

al., 2000). However, a much larger proportion of the genes expressed in the late G2/M phase of

the cell cycle were shown to require ySWI/SNF. In addition, ySWI/SNF was shown to recruit the

HAT Gcn5 to the promoters of many genes in late mitosis (Krebs et al., 2000). An alternative

explanation for the finding that not many genes actually require ySWI/SNF is functional

redundancy with other factors, involving either other ATP-dependent remodeling factors or

histone modifying complexes. In contrast to all currently known genes encoding for ySWI/SNF

components, most of those coding for subunits of RSC are essential, including the gene coding

for its ATPase subunit. RSC is much more abundant than ySWI/SNF. In a search for RSC target

genes, it was found that temperature-sensitive mutant of RSC leads to G2/M cell arrest, and that

ribosomal and cell wall proteins depend on RSC for their expression, which might provide an

explanation for the essential nature ofRSC (Angus-Hill et al., 2001).

In higher eukaryotes, expression of a dominant-negative BRM protein variant in D.

melanogaster caused defects of the peripheral nervous system, homeotic transformations, and

decreased viability (reviewed in (Becker and Hörz, 2002)). Of the two SWI/SNF human

homologues, BRM is dispensable in knockout mice, whereas mouse embryos homozygotes for a

null mutation in BRG1 die early in development, and BRG1 heterozygotes are predisposed to

tumours (Bultman et al., 2000). Recently, microarray based expression profiling identified over

80 genes as target of BRG1 regulation (Liu et al., 2001), identifying, together with additional

studies, BRG1 as a central regulator of the cell cycle and a tumor suppressor (reviewed in

(Becker and Hörz, 2002)).

1.2.2.2. The ISWI Group

This group includes drosophila ISWI, NURF, CHRAC, ACF, yeast ISWI, ISW2 and

human RSF. All of these contain an ATPase subunit that is related to drosophila ISWI. Although

the ISWI group hydrolyses very similar amount ofATP (-100 ATP/min.) as the SWI/SNF group

of enzymes, it has optimal ATPase activity only in the presence of nucleosomal DNA (for review,

see (Langst and Becker, 2001b)). Some features of the remodeled nucleosomes in vitro are listed

in Tab. 1-2.

In drosophila, homozygous null mutation of the ISWI gene is lethal, although

development can proceed until the late larval stages, owing the stockpiles of maternal factor in

the developing embryo (Deuring et al., 2000). Visualisation of the protein on polytene

11

1. Introduction

chromosomes shows that the bulk of ISWI does not correlate with the bulk of RNA polymerase

II, which indicates that transcriptional regulation is unlikely to be the main role of ISWI, at least

in the specialized salivary gland tissue (Deuring et al., 2000). As for SWI/SNF complexes, ISWI-

containing complexes can be directed to specific target sites via transcriptional activators (Xiao

et al., 2001).

In yeast, the two proteins most closely related to ISWI, ISWI and ISW2, also reside in

complexes with nucleosome remodeling and spacing activity (Gelbart et al., 2001; Tsukiyama et

al., 1999). In vitro studies have shown that yISW2 enhances the regular spacing of nucleosomes

within arrays in an ATP-dependent manner (Gelbart et al., 2001; Tsukiyama et al., 1999).

However, in contrast to ylSWl, no nucleosome disruption activity could be detected after

remodeling with yISW2 (Tsukiyama et al., 1999).

In vivo, targeting of the ISW2 complex leads to repression of a set of meiotic genes,

apparently through mobilization of nucleosomes into repressive positions (Goldmark et al.,

2000). ISW2 recruitment has been shown to depend on the transcription factor Ume6 in some

cases and in others not. Chromatin analyses indicate that ISW2 generates repressive structures,

inaccessible to DNasel, at target promoters, presumably through positioning or placement of

nucleosomes (Fazzio et al., 2001; Kent et al., 2001). Interestingly, yISW2 and ySWI/SNF have

both been shown to regulate the expression of the INOI gene, yISW2 being required for

repression and ySWI/SNF for activation of transcription (Fazzio et al., 2001; Goldmark et al.,

2000; Sudarsanam and Winston, 2000; Sugiyama and Nikawa, 2001).

12

Table

1-1:

TheSWI/SNFgroup

Complex

SWI2-likeSubunit

BiochemicalPr

oper

ties

ofRemodeledNucleosomes

SWI/SNF

Swi2/Snf2

S.cerevisiae

RSC

Sthl

S.cerevisiae

INO80

Brahma

Ino80

S.cerevisiae

Brm

Drosophila

•lossoftherotationalph

asin

goftheDNAonthesurfaceofthehistoneoctamer,al

thou

ghtheDNA

remains

atleastpa

rtly

associatedwiththehistoneoctamer(C

ote

et

al.,

1998

)•increasedaccessibilityofthenucleosomalDNA

totranscript

ionfactorsand

restrictionenzymes(Boyer

et

al.,

2000a;

Cote

et

al.,

1998;Jaskelioffetal,2000;Ut

ley

etal,19

97)

•increasedoctamermo

bili

tyinei

s(Jaskelioff

etal,2000;Whitehouse

etal

,19

99)

•increasedsusceptibility

tooctamerdi

spla

ceme

ntintrans(Whitehouse

etal

,1999)

•reductionoftheto

talamountofDNA

associatedwiththehistoneoctamer,asmeasuredbyelectronsp

ectr

osco

pic

imag

ing,

sugg

esti

ngthatDNA

attheedgeofthenucleosome

'pee

lsoff'thehistoneoctamer(B

azet

t-Jo

nes

etal

,19

99)

•

apparentassociationofdisruptednucleosomecoresintoadinucleosome-likesp

ecie

s,th

roug

hinteractionsofloosened

DNA

withhistoneoctamersfromothernucleosomes(S

engu

pta

etal

,2001)

•theevictionorgrossrearrangementofH2A/H2B

dimers

isnotre

quir

edforre

mode

ling

,thesplittingofH3/H4tetramer

neither(B

azet

t-Jo

nes

etal,19

99;Boyer

etal,2000b)

•afterre

mode

ling

,thecontactsbetweencomplexandDNA

arechanged(S

engu

pta

etal

,2001)

•changes

inDNA

topo

logy

arere

quir

edforremodeling

(Gavin

etal

,2001)

•extrudescruciformDNA

fromnakedDNA

andchromatinsubstrates(Havas

etal

,2000)

•HAT

stabilizescomplexbi

ndin

gandenhanceremodeling

(Hassan

etal

,2001)

•lossoftherotationalph

asin

goftheDNAonthesurfaceofthehistoneoctamer,al

thou

ghtheDNA

remains

atleastpa

rtly

associatedwiththehistoneoctamer(C

airn

setal,19

96)

•increasedsusceptibility

tooctamerdi

spla

ceme

ntintrans(L

orch

etal

,19

99)

•

apparentassociationofdisruptednucleosomecoresintoadinucleosome-likesp

ecie

s,th

roug

hinteractionsofloosened

DNA

withhistoneoctamersfromothernucleosomes(L

orch

etal

,1998;Lorch

etal

,2001)

•reductionofnucleosome

stab

ilit

yatelevatedionicstrength

(Lor

chetal

,19

98)

•afterre

mode

ling

,thecontactsbetweencomplexandDNA

arechanged(S

engu

pta

etal

,2001)

•increasedaccessibilityofthenucleosomalDNA

totranscript

ionfactorsandrestrictionenzymes(Shen

etal

,2000)

•RvblandRvb2,twosubunitswhichsharehomology

tothebacterialHo

llid

ayju

ncti

onhelicaseRuvB,

functioninstrand

disp

lace

ment

assaysfor3'to5'helicaseac

tivi

ty(S

hen

etal,2000)

•weaknucleosomesp

acin

gab

ilit

yincomparisonwithISWI(K

aietal

,2000)

•essentialcofactorforZestemediatedtranscriptionfromchromatinte

mpla

tes(K

aietal

,2000)

Table

1-1:

TheSWI/SNFgroup

Complex

SWI2-likeSubunit

BiochemicalPr

oper

ties

ofRemodeledNucleosomes

hSWI/SNF

Brgl

orhBrm

•lossoftherotationalph

asin

goftheDNAonthesurfaceofthehistoneoctamer,al

thou

ghtheDNAremains

atleastpa

rtly

Human

associatedwiththehistoneoctamer(Kwon

etal,19

94)

•increasedaccessibilityofthenucleosomalDNA

totranscript

ionfactorsand

restrictionenzymes(Boyer

etal

,2000a;

Kwon

etal,19

94;Schnitzleretal,19

98)

•increasedsusceptibility

tooctamerdi

spla

ceme

ntintrans(P

hela

netal

,2000)

•

apparentassociationofdisruptednucleosomecoresintoadinucleosome-likesp

ecie

s,th

roug

hinteractionsofloosened

DNA

withhistoneoctamersfromothernucleosomes(P

hela

netal

,2000;

Schnitzleretal

,19

98;Schnitzleretal

,2001)

•relocationofthehistoneH2A

N-terminal

tail

fromaposition

appr

oxim

atel

y40bp

sontheothersideofthenucleosome

dyad

toalocationnearthenucleosomedyad(L

eeetal,19

99)

•thehistoneN-terminal

tailsarenotre

quir

edforremodeling

(Guyon

etal

,19

99)

•extrudecruciformDNA

fromnakedDNAandchromatinsubstrates(Havas

etal

,2000)

Table

1-2:

TheISWIgroup

Complex

ISWI-likeSubunit

BiochemicalPr

oper

ties

ofRemodeledNucleosomes

ISWI

Iswl

•increasedaccessibilityofthenucleosomalDNA

totranscri

ptio

nfactorsandrestrictionenzymes(Tsukiyama

etal

,19

99)

S.cerevisiae

•enhancethere

gula

rsp

acin

gofnucleosomeswithinarrays(Tsukiyama

etal

,19

99)

ISW2

Isw2

•enhancethere

gula

rsp

acin

gofnucleosomeswithinarrays(G

elba

rtetal

,2001

;Tsukiyama

etal

,19

99)

S.cerevisiae

Table

1-2:

TheISWIgroup

Complex

ISWI-likeSubunit

BiochemicalPr

oper

ties

ofRemodeledNucleosomes

NURF

ISWI

Drosophila

ACF

ISWI

Drosophila

dCHRAC

ISWI

Drosophila

RSF

hSnf2h

Human

hACF/hCHRAC

hSnßh

Human

•changes

intheDNaseldigestionpatternofmononucleosomes,ge

nera

ting

new

prot

ecti

onsandenhancements

(TsukiyamaandWu,

1995

)•increasedaccessibilityofthenucleosomalDNA

totranscript

ionfactorsandrestrictionenzymes(Boyer

etal

,2000a)

•increasedoctamermo

bili

tyinei

s(Hamiche

etal,19

99)

•oneormorehistoneN-terminal

tail

isre

quir

edforremodeling(G

eorg

eletal

,1997;Hamiche

etal

,2001)

•noevictionofhistone(Hamiche

etal,19

99)

•de

grad

estheregularsp

acin

gofnucleosomalarrays(V

arga

-Wei

szetal

,19

97)

•increasedaccessibilityofthenucleosomalDNA

totranscript

ionfactors

(Ito

etal

,19

97)

•increasedoctamermo

bili

tyinei

s(E

berh

arte

retal,2001)

•enhancestheregularsp

acin

gofnucleosomeswithinarraysor

facilitatesnucleosomeassembly

(Ito

etal

,1997;

Nakagawa

etal,2001)

•enhanced

stab

ilit

yofmononucleosomes

toMNase

digestion(L

angs

tetal

,19

99)

•increasedaccessibilityofthenucleosomalDNA

totranscript

ionfactorsandrestrictionenzymes(Boyer

etal

,2000a;

Varga-Weisz

etal,19

97)

•increasedoctamermo

bili

tyinei

s(E

berh

arte

retal,2001;Langst

etal

,19

99)

•oneormorehistoneN-terminal

tail

isre

quir

edforremodeling

(Cla

pier

etal

,2001)

•enhancestheregularsp

acin

gofnucleosomeswithinarraysorfacilitatesnucleosomeassembly(V

arga

-Wei

szetal

,19

97)

•bindstonucleosomeson

lyifth

eycontainover147bpsofDNA(Brehm

etal

,2000)

•enhancestheregularsp

acin

gofnucleosomeswithinarrays(LeRoy

etal

,1998)

•directsVP16-mediatedtranscriptionfromchromatinte

mplate

s(LeRoy

etal

,19

98)

•enhancestheregularsp

acin

gofnucleosomeswithinarraysorfacilitatesnucleosomeassembly(LeRoy

etal

,2000)

1 Introduction

1.2.2.3. The Mi-2 Group

This very recently discovered group includes human NURD, xenopus Mi-2 and drosophila

Mi-2. They contain an ATPase subunit highly related to the human Mi-2 and show, in addition to

chromatin-remodeling, also histone deacetylase activities (Tong et al, 1998; Wade et al, 1998;

Xue et al, 1998; Zhang et al, 1998b). The Mi-2 complexes are believed to repress transcription

through their remodeling and deacetylation activities in a targeted manner. Targeting might occur

via direct interaction with sequence-specific transcriptional repressors (Kehle et al, 1998; Kim et

al, 1999; Xue et al, 1998). Alternatively, Mi-2 complexes might be targeted to methylated DNA

(Wade et al, 1999; Zhang et al, 1999). Recently, a novel Mi-2-type ATPase, Hrpl, has been

discovered in fission yeast. The phenotype of a hrpl deletion points to an involvement of the

enzyme in chromosome condensation in mitosis (Yoo et al, 2000).

Recombinant dMi-2 was shown to promote nucleosome mobilisation in an ATP-dependent

manner in vitro, irrespective of the presence or absence of H4 N-terminal tails (Brehm et al,

2000).

1.3. UV Damage Formation

The ultraviolet (UV) component of solar radiation is a major source of physical damage in

DNA. The UV radiation spectrum has been subdivided into three wavelength bands designated

UV-A (400 to 320 nm), UV-B (320 to 290 nm) and UV-C (290 to 100 nm). Solar UV radiation

consists mainly of UV-A and UV-B, since wavelength up to 300 nm are absorbed by the

stratospheric ozone layer. Formation of photolesions is highest at 254 nm, which is close to the

absorption maximum of the DNA bases. Identical lesions are induced at longer wavelength,

although at lower rates (see (Friedberg et al, 1995)).

1.3.1. UV-Photoproducts

Most UV lesions are formed at adjacent pyrimidines. Pyrimidine clusters are hot spots for

UV induced damages and mutations (Miller, 1985). The two major classes of stable UV

photoproducts are cyclobutane pyrimidine dimers (CPDs) and pyrimidine (6-4) pyrimidone

photoproducts (6-4)PPs (Fig. 1-1). CPDs are the most abundant lesions, being induced four to

five times more frequently than (6-4)PPs at most sites (Mitchell and Nairn, 1989). Besides CPDs

and (6-4)PPs, a number of rare other lesions were identified, which account for less than 1% of

total UV lesions and include spore photoproducts, purine dimers, pyrimidine hydrates, thymine

glycols or DNA-protein crosslinks ((Sage, 1993), reviewed in (Friedberg et al, 1995)).

16

1. Introduction

A.

i rrcH>

N >0

c.

cis,syn-

cyclobutane pyrimidine dimer

CPD

5'

Io

.

Io-p.

O-f »O \

t n

Ay""/" o

pyrimidine (6-4) pyrimidone

photoproduct

(6-4)PP

Figure 1-1: The two major photoproducts generated by UV light.A. Cyclobutane pyrimidine dimers are the predominant UV photoproduct. Formation

occurs between any two adjacent pyrimidines but most frequently as thymine-thymineCPD. B. CPDs introduce a bend of 7 to 9° in the DNA double helix. C. (6-4)

pyrimidine-pyrimidone photoproduct is the second major UV photoproduct and is

predominantly formed at thymine-cytosine sites. D. (6-4)PPs induce a bend of about

44° in the DNA double helix. Adapted from (Doetsch, 1995) and (Kim et al, 1995).

CPDs are formed when two adjacent pyrimidines are covalently crosslinked by the

formation of a cyclobutane ring between their 5, 6 double bonds. Out of its 12 possible isomeric

forms, the CPD exists predominantly in the cis-syn form in double-stranded B-form DNA. NMR

and gel retardation studies revealed that a cis-syn CPD induces a 7 to 9° bend in duplex DNA

(Kim et al, 1995; Wang and Taylor, 1991). The yield of CPD formation is dependent on the

sequence: CPDs are preferentially induced at TT and significantly less at C-containing sites

(Setlow, 1968; Setlow and Carrier, 1966). Irradiating plasmid DNA at 254 nm, a ratio of

TT:CT:TC:CC (nucleotides in 5' to 3' direction) of 68:13:16:3 was formed (Mitchell et al,

1992). In addition, the identity of the bases flanking the potential dimer sites influences CPD

formation. Presence of a pyrimidine 5' to a dipyrimidine enhanced the potential for dimerisation

(Bourre et al, 1987; Gordon and Haseltine, 1980). Furthermore, the dimer formation was

inhibited to a greater extent by a 5' flanking guanine than by an adenine (Mitchell et al, 1992).

17

1 Introduction

(6-4)PPs are formed when two adjacent pyrimidines are linked by a covalent single bond

between positions 6 and 4. NMR studies and structural calculation established that (6-4)PPs lead

to a bending of 44° in the DNA double helix ofB-DNA (Kim and Choi, 1995; Kim et al, 1995).

(6-4)PP are formed preferentially at TC sites, followed by CC, and to a lesser extent at TT,

whereas CT sites seem refractory to (6-4)PP formation (Lippke et al, 1981; Mitchell and Nairn,

1989). Irradiation of (6-4)PPs with 313 nm light leads to the formation of the Dewar isomer

(Taylor et al, 1988), which is relevant for sunlight carcinogenesis (Mitchell and Nairn, 1989).

From experiments in mammalian cells, the prevailing view considers the (6-4) PPs much more

mutagenic than the CPDs (e.g. (Gentil et al, 1996; Gentil et al, 1997; Zdzienicka et al, 1992)).

Different structural distortions caused by (6-4)PPs and CPDs may explain the large differences in

mutation spectrum and repair activities.

1.3.2. UV Damage Formation

The torsional flexibility of the DNA helix at the site of the photoproduct plays a

significant role in determining the ability of a given base pair to form a dimer, since extensive

rotation of the neighbouring pyrimidines from their usual B-form DNA conformation is required

for the formation of CPDs and (6-4)PPs (Becker and Wang, 1989b).

DNA sequence that are easily unwound or bent are favourable sites for damage formation,

whereas rigid DNA structures interfere with photodimerisation (Becker and Wang, 1989a;

Lyamichev et al, 1990; Tang et al, 1991). For example, CPDs form at higher yields in single-

stranded DNA (Becker and Wang, 1989b) and at the flexible ends of poly(dA)(dT) tracts, but not

in their rigid centre (Lyamichev, 1991). Protein-DNA contacts often cause distortions in the

regular B-DNA structure, thereby affecting both yields and types ofUV damage formation (UV

footprinting; e.g. (Aboussekhra and Thoma, 1999; Becker and Wang, 1984; Pfeifer et al, 1992;

Seileck and Majors, 1987; Seileck and Majors, 1988)). UV-photofootprint of the EcoRI-

endonuclease DNA complex revealed that kinking of the DNA by EcoRI greatly enhances the

UV photoreactivity of DNA at the site of the kink, but contacts between the endonuclease and

the DNA bases have an inhibitory effect on UV-damage formation (Becker et al, 1988). This

example illustrate the difficulty to predict how protein-induced conformational changes in DNA

will influence photoproducts yields.

Chromatin structure affects UV damage formation due to the distortion of DNA on the

histone octamer. Most of the structural data concerning DNA damage formation in nucleosomes

were obtained from mixed sequence populations. Formation of CPDs in mixed sequence

nucleosome cores is significantly modulated with peaks at intervals of 10.3 bps (Gale et al.

18

1. Introduction

1987; Gale and Smerdon, 1988). Maxima were observed at positions farthest from the histone

surface very similar to the cutting pattern of DNasel. Two observations indicate that the

modulation of CPD formation in nucleosomes is determined by DNA curvature rather than

histone-DNA contacts. (1) In a DNA loop free of protein contacts, the relation of the CPD peaks

to the curvature of the loop was similar to that in the nucleosome (Pehrson and Cohen, 1992). (2)

Loss of periodicity was observed in unfolded nucleosomes (Brown et al, 1993).

The effects of individual nucleosomes on UV photoproduct formation are different. The

yields and distribution of CPDs on a nucleosome reconstituted on a define sequence (HISAT)

resemble only partially those of mixed sequence nucleosomes (Schieferstein and Thoma, 1996).

Thus, the observed modulation in mixed sequence displays an average damage formation

pattern. On the other hand, the damage formation pattern of individual nucleosomes reflects the

particular structural features of these nucleosomes.

CPDs and (6-4)PPs are detected throughout the nucleosome core, suggesting that UV

lesion can be tolerated even at positions where the damage-induced distortions do not coincide

with the natural distortions of nucleosomal DNA. On the other hand, folding of DNA in

nucleosomes exerts structural constraints that can disrupt the rigid structure of T-tracts (Hayes et

al, 1991; Schieferstein and Thoma, 1996). Thus, nucleosomes reveal substantial degree ofDNA

flexibility as well as structural constraints, which modulate damage formation and might as well

be essential for damage recognition.

CPDs and (6-4)PPs distort the DNA helix and may thereby alter the histone-DNA

interactions, since the orientation of DNA in nucleosomes is influenced by its sequence-

dependent structure and flexibility (Simpson, 1991). The stability of nucleosomes was altered

after UV irradiation of plasmid DNA assembled with nucleosomes at 3000 J/m2 in vitro

(Matsumoto et al, 1994). Nucleosome reconstitution was shown to be less efficient on UV

irradiated DNA and to lead to alternate positions of the histone octamer compared with

reconstitutions on non-damaged DNA (Mann et al, 1997; Matsumoto et al, 1994; Schieferstein

and Thoma, 1996). However, irradiation with UV light did not alter neither the rotational nor the

translational setting of pre-assembled nucleosomes (Liu et al, 2000; Schieferstein and Thoma,

1996).

19

1. Introduction

1.4. Repair of UV Lesions

In most organisms, three basic mechanisms operate to remove CPDs and (6-4)PPs from

DNA: the nucleotide excision repair (NER) pathway, which also removes other bulky DNA

lesions (de Laat et al, 1999); direct reversal by DNA photolyases (photoreactivation; (Sancar,

2000)); and base excision by UV damage specific endonucleases (Lloyd, 1999).

1.4.1. Nucleotide Excision Repair

Nucleotide excision repair (NER) is a repair pathway which is conserved from prokaryotes

to humans. NER is a multistep mechanism involving about 30 proteins, which remove a wide

class ofDNA double helix distorting lesions, including CPDs and (6-4) PPs. NER is not essential

for cell viability, but defects in repair genes cause the sun-sensitive, cancer-prone genetic

disorders xeroderma pigmentosum (XP), Cokayne's syndrom (CS) and trichothiodystrophy

(TTD) (reviewed in (Hoeijmakers, 2001a)). The basic mechanism of NER involves five steps:

DNA damage recognition, incision at 5' and 3' side of the lesion by the action of two

endonucleases, removal of the excised oligonucleotide, gap filling by repair synthesis using the

complementary strand as a template and sealing the patch by DNA ligases (reviewed in (de Laat

et al, 1999; Prakash and Prakash, 2000)).

NER is divided in two sub-pathways, global-genome repair (GG-NER) which refers to

repair of the genome overall, and transcription-coupled repair (TC-NER) which leads to

enhanced removal of lesions on the transcribed strand of active genes (reviewed in

(Hoeijmakers, 2001a; Tornaletti and Hanawalt, 1999)). In TC-NER, RNA polymerase II stalled

at a DNA lesion may serve as damage sensor and promote NER (reviewed in (Citterio et al,

2000b; de Laat et al, 1999)).

1.4.2. Photoreactivation

As an alternative or additional pathway to NER, a wide variety of organisms including

bacteria, fungi, plants, invertebrates and many vertebrates can specifically revert photoproducts

to their native bases by DNA photolyase (photoreactivation) (Sancar, 2000). No photolyase was

found in humans (Sancar, 1996; Todo, 1999; Yasui et al, 1994). Photoreactivation is a two-step

mechanism allowing the direct repair of both CPDs and (6-4)PPs. First, the UV lesion is

recognized in a light independent way (dark reaction). Second, the energy of photoreactivating

blue light is used to split the covalently-linked pyrimidines (enzyme-catalyzed monomerization).

20

1. Introduction

This light reaction is catalyzed by two non-covalently attached prosthetic groups, one ofwhich is

the catalytic cofactor 1,5-dihydroflavin adenine dinucleotide (FADH"), and the other a light

harvesting cofactor.

Photolyases are monomeric proteins of 50-60 kDa which belong to the family of DNA

photolyase/blue light photoreceptor. This protein family consists of three groups: CPD

photolyases and (6-4)PP photolyases, which specifically remove CPDs and (6-4)PPs,

respectively, and blue light photoreceptors (cryptochromes), which function in signal

transduction (reviewed in (Kanai et al, 1997; Todo, 1999)).

1.4.2.1. CPD Photolyases

The CPD photolyase gene was first cloned in E. coli (Sancar and Rupert, 1978). Based on

sequence homology, CPD photolyases are subdivided into class I and class II CPD photolyases

with less than 20% sequence identity across the class boundary (Yasui et al, 1994). Class I CPD

photolyases were isolated from pro- and eukaryotic unicellular organisms and are further divided

into two groups according to the chemical nature of their light-harvesting chromophore: 5,10-

methenyltetrahydrofolate (MTHF) or 8-hydroxy-5-deazariboflavin (8-HDF), with absorption

maxima of-380 nm and -440 nm, respectively (Johnson et al, 1988; Sancar et al, 1987b). Both

the E. coli and the S. cerevisiae photolyase are MTHF class I enzymes. Class II photolyases were

found in eubacteria, archaebacteria and higher eukaryotes, e.g. in goldfish C. auratus (Yasuhira

and Yasui, 1992), D. melanogaster (Todo et al, 1994) and the marsupial M. domestica (Kato et

al, 1994). Besides these multicellular organisms, class II photolyases were also isolated from

archaebacteria (Kiener et al, 1989; Yasui et al, 1994) and eubacteria (O'Connor et al, 1996).

Recently, a CPD class II photolyase that complements a photolyase-deficient E. coli strain,

was found in fowlpox virus (Srinivasan et al, 2001). Furthermore, sequencing ofthe genomes of

an entomopox virus, a rabbit obroma virus and myxoma virus revealed putative class II

photolyase genes, suggesting that enzymatic photoreactivation might be important for viruses

(Afonso et al, 1999; Afonso et al, 2000; Cameron et al, 1999; Willer et al, 1999).

In yeast, there are about 250 to 300 molecules of photolyase per cell under constitutive

conditions (Yasui and Laskowski, 1975). Photolyase expression is induced by DNA damaging

agents (reviewed in Sancar (Sancar, 2000)). 5- to 10-fold induction was observed after treatment

with UV light or alkylating agents and up to two-fold induction was measured after bleomycin

treatment or heat shock at 37°C (Sebastian et al, 1990). Thus, it seems likely that a global

damage response pathway regulates photolyase expression.

21

1. Introduction

The amino acid sequence of S. cerevisiae photolyase displays a striking homology to E.

coli photolyase, particularly in the carboxy-terminal 150 amino acids, where the sequence

identity is >70% (Sancar, 1985b; Yasui and Langeveld, 1985). The conserved carboxy-terminus

comprises the flavin binding domain as well as several of the amino acids that are predicted to

contact DNA (Baer and Sancar, 1993; Malhotra et al, 1992; Park et al, 1995; Vande Berg and

Sancar, 1998). Both S. cerevisiae and E. coli photolyases contain MTHF as light-harvesting

cofactor (Johnson et al, 1988; Sancar et al, 1984; Sancar et al, 1987b) and protect a 6-8

nucleotide region surrounding the dimer from chemical attack on the dimer-containing strand

(Baer and Sancar, 1989; Husain and Sancar, 1987). In addition, S. cerevisiae and E. coli

photolyase genes can partially cross-complement each other in photoreactivation deficient

mutants of both strains ((Langeveld et al, 1985; Sancar, 1985a) and are thought to repair CDPs

via a 'flip-out' mechanism (see Section 1.4.2.3.; (Park et al, 1995; Vande Berg and Sancar,

1998)).

1.4.2.2. (6-4)PPPhotolyases

(6-4)PP specific photolyases were cloned and characterised from D. melanogaster (Kim et

al, 1996a; Todo et al, 1996), X. laevis (Kim et al, 1996b; Todo et al, 1997) and A. thaliana

(Nakajima et al, 1998). Additionally, (6-4)PP photolyase activity was detected in cultured

goldfish cells (Uchida et al, 1997) and a (6-4)PP repair activity was suggested for the phrA gene

product in E. Coli (Dorrell et al, 1993; Dorrell et al, 1995). D. melanogaster (6-4)PP photolyase

gene exhibits sequence similarity to CPD class I photolyases (Todo et al, 1996). The drosophila

(6-4)PP photolyase restores (6-4)PPs to canonical bases, as does CPD photolyase with CPDs.

However, the efficiency of repair per incident photon is 100-fold lower compared to CPD

photolyases (Kim et al, 1994).

1.4.2.3. The Molecular Mechanism ofPhotoreactivation

The molecular mechanism of photoreactivation has been extensively studied in E. coli

(Sancar, 1994; Sancar et al, 1987a). The light harvesting chromophore MTHF absorbs a photon

and transfer the excitation energy to the catalytic cofactor FADH". The enzyme transfers an

electron from FADH to the CPD to catalyse its fission yielding the two original pyrimidine. After

the reversal of the dimer, the electron is transferred back to the FADH, regenerating catalytically

active FADH" (reviewed in (Hearst, 1995)). Studies using time-resolved absorption spectroscopy

suggests intraprotein radical transfer from excited FADH" to the dimer via several tryptophan

22

1. Introduction

close to the catalytic cofactor (Aubert et al, 2000). Similar mechanisms exist in class II CPD

photolyases (Aubert et al, 1999) and (6-4) PP photolyases (Hitomi et al, 1997; Zhao et al,

1997).

Detailed insight into the photoreactivation process was obtained from the crystal structure

of the CPD photolyases from E. coli (Park et al, 1995; Park et al, 1993) and A. nidulans ((Miki

et al, 1993; Tamada et al, 1997); reviewed in (Deisenhofer, 2000)). E. coli photolyase is of the

MTHF type, while A. nidulans photolyase contains 8- HDF as the light-harvesting cofactor. The

two structures are very similar. The FAD cofactor is accessible through a hole in the surface of

the protein. Dimensions and polarity of the hole match those of a pyrimidine dinucleotide,

suggesting that the dimer bases 'flip out' of the helix to fit into this pocket. This would allow the

direct contact of the UV lesion and the catalytic cofactor. Although high-quality co-crystals of

photolyase with its substrate are not yet available, modelling of the enzyme-substrate complexes

of S. cerevisiae photolyase substantiates the 'dinucleotide flipping' model (Vande Berg and

Sancar, 1998).

Base flipping occurs in many systems where enzymes need access to a DNA base to

perform chemistry on it. It has been directly observed for the cytosine-5 methyltransferase

M.Hhal (Klimasauskas et al, 1994) and M.HaeIII (Reinisch et al, 1995), and for the repair

enzymes T4endonucleaseV (Vassylyev et al, 1995) and human uracil DNA glycosylase

(Slupphaug et al, 1996). In addition, crystal structures of many proteins, including several DNA

methyltransferases, nucleases and glycosylases, suggest the use of similar base flipping

mechanisms (reviewed in (Roberts and Cheng, 1998)).

1.4.3. CPD Glycosylases

Another UV damage specific enzyme is CPD glycosylase, which introduces a nick at one

of the glycosyl bonds in a CPD and generates an abasic site. The resulting abasic site is a

substrate for base excision repair (BER). CPD glycosylase is found in Microccocus luteus

(Haseltine et al, 1980) and bacteriophage T4-infected E. coli (T4endonucleaseV; (Radany and

Friedberg, 1980)). Recently, additional CPD glycosylases were found in a Chlorella virus

(Furuta et al, 1997; Garvish and Lloyd, 1999) and in the bacteria Neisseria mucosa (Nyaga and

Lloyd, 2000) and Bacillus sphaericus (Vasquez et al, 2000).

T4endonucleaseV (T4endoV) is a DNA glycosylase/AP lyase that can initiate repair of

cis-syn CPDs in DNA by cleaving the glycosydic bond of the 5' pyrimidine and then cleaving

the phosphodiester backbone (reviewed in (Lloyd, 1999)). It has been shown to restore repair of

UV damage in excision-repair deficient cells in E. coli (Chenevert et al, 1986; Valerie et al.

23

1. Introduction

1985) and in many eukaryotes including yeast, drosophila and human (Arrand et al, 1987;

Banga et al, 1989; Chenevert et al, 1986; Valerie et al, 1986). Since the excision-repair defects

(radl, rad2, rad3, rad4 and radlO mutants of S. cerevisiae and human Xeroderma D cells)

involved a failure to initiate incision at sites of DNA damage, T4endoV appears to mimic the

incision step ofNER (reviewed in (Memisoglu and Samson, 1996)).

1.5. Repair in Chromatin

The histone octamer exerts a dominant constraint on the structure of DNA in the

nucleosome, thereby affecting damage accessibility and repair. NER, photoreactivation and CPD

glycosylase incision are all modulated by chromatin structure (reviewed in (Smerdon and

Conconi, 1999; Thoma, 1999)).

1.5.1. NER and Chromatin

The complexity of the NER pathway makes it difficult to imagine how DNA lesions can

be recognized and processed in chromatin. Chromatin may interfere with NER factors at the

level of damage recognition or/and repair processing. In the reconstituted yeast NER system,

damage recognition is supported by the Rad7/Radl6 complex, which, together with the Rad4/

Rad23 complex, binds to UV damaged DNA synergistically and in an ATP-dependent manner

(Guzder et al, 1997; Guzder et al, 1999). It is not known whether the damage recognition

complexes can interact with lesions on the nucleosome surface. In the open repair complex,

about 25 bps ofDNA are unwound (Evans et al, 1997), and the human excision repair complex

requires -100 bps of DNA to excise the lesion in vitro (Huang and Sancar, 1994). Such a

complex appears to be incompatible with the structure of nucleosomes. Moreover, the linker

DNA between nucleosomes (0-90 bps) is too short to accommodate a repair complex. Hence,

rearrangement or (partial) unfolding of nucleosomes are likely to be required for NER in

chromatin.

Using crude cell extracts to perform NER in vitro, it was shown that repair synthesis is

strongly reduced on UV-irradiated DNA pre-assembled in nucleosomes (Wang et al, 1991) or

UV-irradiated simian virus 40 (SV40) minichromosomes (Sugasawa et al, 1993), compared to

naked DNA. A study of repair at specific sites in a mononucleosome demonstrated that CPD

removal is inhibited at most nucleosomal positions and is not influenced by rotational setting

24

1 Introduction

(Liu and Smerdon, 2000). On the other hand, Xenopus extract proficient for NER can repair a

lesion placed in reconstituted nucleosomes (Kosmoski et al, 2001), suggesting that factors

responsible for modulating chromatin structure may be present in the extracts.

The efficiency of damage excision carried out by purified NER factors is lower on both

UV-irradiated SV40 minichromosomes (after long incubation times) and reconstituted

mononucleosomes than on naked DNA (Araki et al, 2000; Hara et al, 2000). Recently, the use

of a dinucleosomal substrate demonstrated that excision of (6-4)PPs by purified factors is

strongly inhibited even when the lesion is located within the linker DNA. However, in the

presence of the chromatin assembly and remodeling factor ACF and ATP, repair was enhanced in

the linker but not in nucleosomes (Ura et al, 2001).

In contrast to the severe repair inhibition observed on the nucleosome surface in vitro,

NER is complete in vivo (reviewed in (Smerdon and Conconi, 1999; Thoma, 1999)). To gain

information about NER in living cells, removal of UV-lesions was compared with chromatin

structure in yeast minichromosomes. Modulations of DNA repair were observed and correlated

with gene expression as well as with nucleosome positions and stability (Smerdon et al, 1990;

Smerdon and Thoma, 1990). CPDs and (6-4)PPs were mapped at high resolution by primer

extension with Taq polymerase, which is blocked at CPDs and (6-4)PPs (Wellinger and Thoma,

1996), in a yeast minichromosome containing the active URA3 gene. While repair rates on the

transcribed strand were dominated by transcription-coupled repair and showed no correlation

with chromatin structure, analysis of the non-transcribed strand revealed pronounced

heterogeneity in repair rates. Fast repair correlated with lesions located in linker DNA and

toward the 5' end of a nucleosome. Slow repair correlated with the nucleosomes positions

(Tanaka et al, 1996; Wellinger and Thoma, 1997). Similar results were reported for the genomic

copy of the URA3 gene and for removal of (6-4)PPs, indicating that modulation by chromatin

structure is not damage dependent (Tijsterman et al, 1999). These results provide strong

evidence that NER is modulated by the arrangement of nucleosomes along the DNA. In addition,

NER has been shown to be modulated by other protein/DNA interactions (Aboussekhra and

Thoma, 1999; Gao et al, 1994; Suter et al, 2000; Tu et al, 1996).

1.5.2. Photoreactivation in Chromatin

In eukaryotic cells, photolyase has to deal with chromatin as substrate. Chromatin may

interfere with photoreactivation at the level of damage recognition or/and at the level of damage

flip-out. CPD accessibility and repair in nucleosomes were tested in vitro using reconstituted

nucleosomes as model substrates and two damage-specific enzymes, T4endoV and E. coli DNA

25

1. Introduction

photolyase. Although T4endoV and photolyase were very efficient in naked DNA, their activity

was dramatically reduced on the surface of the reconstituted nucleosomes (Kosmoski and

Smerdon, 1999; Schieferstein and Thoma, 1998). Thus, folding of DNA into nucleosomes

efficiently protects DNA from being repaired. On the HISAT nucleosome (Schieferstein and

Thoma, 1998), site-specific repair common for T4endoV and photolyase was observed,

indicating that the repair inhibition was due to the structure of the nucleosome and not the

individual properties of the repair enzymes.

In contrast to the severe inhibition of photoreactivation on the nucleosome surface in vitro,

repair by photolyase is complete in vivo (reviewed in (Thoma, 1999)). The first indication that

dynamic properties of chromatin could affect a DNA repair process came from the observation

that -75% ofthe DNA was shielded from photorepair immediately after UV exposure in chicken

embryo fibroblasts, but that all sites became available after 9-12 h (Pendrys, 1983).

Direct information on how photolyase interacts with nucleosomes, linker DNA and non-

nucleosomal regions was obtained by comparison of CPD removal with chromatin structures in

NER deficient yeast strains (Suter et al, 1997). In minichromosomes, photolyase is fast in linker

DNA and nuclease-sensitive regions such as promoters, 3' ends of genes or ARS. On the other

hand, lesions which mapped within the footprint of positioned nucleosomes are removed much

slower. Thus, photoreactivation in living cells is tightly modulated by chromatin structure.

High resolution analysis of CPD repair in the six nucleosomes covering the URA3 gene

demonstrated that photorepair is further modulated on the nucleosome surface (Suter et al,

2002). Lesions located around the dyad axis of nucleosomes are repaired slower than those

located towards the edges. Intrinsic properties of nucleosomes, such as octamer mobility along

the DNA, might explain this modulation, consistent with the observation ofmultiple nucleosome

positions in the URA3 gene (Tanaka et al, 1996). Alternatively, dissociation/reassembly or

partial unfolding of nucleosomes could enhance damage accessibility. Since chromatin

remodeling can promote both nucleosome sliding and disruption, it is conceivable that such

activities contribute to damage accessibility in vivo.

1.5.3. Repair and Chromatin Remodeling

In principle, all reactions that involve DNA can be regulated by altering DNA packaging

and hence DNA accessibility. Since chromatin-modifying complexes play a role in regulating the

accessibility of chromatin in the context of transcription, it is conceivable that they might also

play a role in other nuclear processes, including DNA replication, recombination and repair

(reviewed in (Fyodorov and Kadonaga, 2001; Green and Almouzni, 2002; Narlikar et al, 2002)).

26

1. Introduction

Intrinsic dynamic properties of nucleosomes, like octamer mobility, might place a nucleosomal

lesion in the linker DNA, thus providing a window of accessibility for damage recognition (Suter

et al, 2002).

Alternatively, there is an increasing number of repair related proteins with potential roles

in chromatin remodeling (Green and Almouzni, 2002). CSB and its yeast homologue Rad26 both

belong to the SNF2 family (Eisen et al, 1995). CSB mutants are defective in transcription

coupled repair, while maintaining a proficient global genome repair of many DNA-damaging

agents (Evans and Bohr, 1994; Venema et al, 1990). In addition to its role in DNA repair, the

CSB protein is involved in basal transcription (Balajee et al, 1997), and interacts with the

elongating form of RNA polymerase II (Selby and Sancar, 1997; van Gool et al, 1997).

Recombinant CSB was shown to remodel undamaged nucleosomes and nucleosome arrays in

vitro thus being the first repair enzyme with remodeling activity (Citterio et al, 2000a). Thus,

CSB may have a role in chromatin remodeling during transcription elongation and/or DNA

repair. Rad7-Radl6 is a complex ofthe NER pathway ofyeast S. cerevisiae which is essential for

repair of nontranscribed chromatin (Bang et al, 1992; Verhage et al, 1994) and recognises UV-

lesions in an ATP-dependent way in vitro (Guzder et al, 1998). Radio has homology to SNF2

and might play a role in nucleosome remodeling to generate space for the other NER proteins

(Eisen et al, 1995; Thoma, 1999).

Nucleosome remodeling activity was shown for the yeast INO80 complex which, beside

its ATPase, also contains two proteins related to the bacterial RuvB DNA helicase that catalyses

branch migration of Holliday junctions (Jonsson et al, 2001). In yeast cells, mutation in INO80

causes sensitivity to hydroxyurea, methyl methanesulfonate, ultraviolet and ionizing radiations

(Shen et al, 2000), suggesting a function of INO80 in repair. ACF, on the other hand, is a

chromatin assembly and remodeling factor containing ISWI and Acf1, which was reported to

facilitate NER of a specific lesion in linker DNA of a dinucleosome in vitro, but not of a lesion

positioned in the center of one of the nucleosomes (Ura et al, 2001).

The covalent modification of histones may also contribute to facilitate damage site

accessibility. In vitro, histone H3 acetyl-transferase activity can be directed to sites of UV

damage via the TBP-free TAFII complex, which contains both a DNA-damage binding and a

HAT subunit (Brand et al, 2001). Furthermore, the damage-specific DNA binding proteins XP-E

and DDB, which are involved in the initial steps of NER, have been shown to interact with the

27

1. Introduction

p300 and SAGA HAT complexes, respectively (Datta et al, 2001; Martinez et al, 2001).

Recently, thymine DNA glycosylase, an enzyme involved in the initiation step of base excision

repair, has been shown to interact with the CBP/p300 acetylase (Tini et al, 2002).

These findings support the idea that common strategies and factors are shared by the

different DNA transaction pathways to access DNA within chromatin.

1.6. Aim of Project

In contrast to the complete repair of DNA lesions observed in eukaryotic cells,

nucleosomes have been shown to exert a strong inhibition on repair in vitro. It is not known

whether DNA-lesions become accessible by intrinsic dynamic properties of nucleosomes alone

or whether remodeling activities are required in living cells. Previous experiments that

established a repressive role of nucleosomes in photorepair used nucleosomes reconstituted on

short fragments without space for mobility (Schieferstein and Thoma, 1998). Here, we used

nucleosome reconstitution on an extended DNA sequence (ATDED-long) to unravel the roles of

nucleosome mobility and nucleosome remodeling on CPD formation and accessibility to

photolyase.

ATDED-long is a 226 bps DNA fragment originating from the yeast DEDI promoter and

contains T-tracts of various length on both strand which are hot-spots for CPD formation. DNA

fragments containing this sequence have been shown to form positioned nucleosomes (Losa et

al, 1990). Thus, nucleosomes reconstituted on ATDED-long DNA generate a chromatin model

substrate containing both nucleosomal and naked DNA on which CPD damage formation and

repair can be analysed. E.coli photolyase was used as a tool to assess CPD accessibility of flip-

out enzymes to nucleosomal substrates.

We wished to investigate whether the presence of 'linker' DNA, which gives more space

to the positioned ATDED-long nucleosome, would enhance repair ofnucleosomal CPD, possibly

through nucleosome mobility.

It has been postulated that ATP-dependent chromatin remodeling complexes might play a

role in repair of nucleosomal lesions. However, it is not known whether those complexes work

on damaged nucleosome and whether remodeling facilitates damage recognition and repair. To

assess these questions, we tested the activity of two remodeling machines, ySWI/SNF and

yISW2, on UV damaged ATDED-long nucleosomes and their effects on CPD accessibility to

photolyase.

28

2. Results

2. Results

2.1. The ATDED-long Nucleosome

In living cells, CPD repair by photolyase is complete in spite of the packaging of DNA

into chromatin (Suter et al, 1997). However, photoreactivation is severely inhibited in

nucleosomes reconstituted on a 134 bps long DNA fragment (Schieferstein and Thoma, 1998),

suggesting that additional features, such as nucleosome mobility, might be required for repair of

nucleosomal CPDs. In order to generate a chromatin model substrate which would allow

nucleosome sliding, we used a ATDED-long fragment as a template for nucleosome

reconstitution. ATDED-long contains polypyrimidine tracts on both strands (Fig. 2-1) which

allow CPD repair analysis. It was previously shown that nucleosomes reconstituted onto a DNA

fragment containing the ATDED sequence occupy a single translational and rotational setting

(Losa et al, 1990).

Afllll

Pstl Hhal Xhol

, II 2 3 4 1 56 7 89

<Z t ~~2>

1 1 '

io'Ti 12 13 1415 16 1718 19

i i i i i i i i i i i i l 1

MU 1300 MU1400 MU 1500

(1) 5'tcaTTct

(2) S'cTTTccTTTTTTcTTTTT

(3) S'cTTTTTc

(4) 5'TTTTTTTTTctcTT

(5) 5'TT

(6) 5'TT

(7) 5'TTcc

(8) s'TTTc

(9) 5'TT

(10) cTT5'

(11) ctcc5'

(12) cTT5'

(13) ctcTTTTTTTT5'

(14) TTTTctct5'

(15) cctccTT5'

(16) cccTTTTTcaatc5'

(17) ctatcc5'

(18) TTc5'

(19) TTcTTgTTgTTcTTTTc5'

Figure 2-1: Schematic drawing of the ATDED-long nucleosome

Indicated are map units (MU), position of the radioactive end-label (asterisks), pyrimidine clusters on

top and bottom strand (DNA sequences numbered from 1 to 19) and cutting sites of restriction enzymes

Pstl, AflHI, Hhal and Xhol. The ellipse denotes the location of the histone octamer on the DNA

fragment. The position of the nucleosome centre is indicated (black dot).

29

2. Results

2.1.1. Characterization of the ATDED-long Nucleosome

The 226 bps long SmallEcoRI fragments were end-labeled with 32P on either strand and

nucleosomes were reconstituted by histone octamer transfer from chicken erythrocytes core

particles (Schieferstein and Thoma, 1998). The reconstituted nucleosomes were characterized by

native nucleoprotein gel electrophoresis, DNasel footprinting, restriction enzyme cleavage and

UV footprinting. Native nucleoprotein gel electrophoresis showed that reconstitution was

efficient, with over 90% of the labeled DNA folded into nucleosomes (Fig. 2-2).

Z a Z a

O fc Ö S5

Figure 2-2: Native nucleoprotein gel electrophoresisResults of the top and bottom strands are shown. ATDED-

long DNA (DNA) was reconstituted into nucleosomes

(Nucl). Bands represent naked DNA (D) and nucleosomes

NW

D

Top Bottom

In nucleosomal DNA, DNasel cuts where the minor groove faces away from the histone

surface, resulting in a 10 base repeat pattern on denaturing gels (Lutter, 1978). DNasel digests of

the reconstituted nucleosomes produced the stereotypical 10 bps repeat pattern with either strand

labeled (Fig. 2-3 and Fig. 2-4, lanes 3 to 6, respectively) while naked DNA digests exhibited no

such pattern (lanes 7 to 9). The 10 bps repeat pattern spans the 140 bps between MU 1360 and

1497 (black dots), suggesting that nucleosomes assume a single rotational settings toward the

end of the fragment, with the dyad axis close to MU 1430. Furthermore, the nucleosome-specific

cutting sites on both strands correlated.

Additional DNasel sensitive sites were observed around MU 1378 and 1469 with either

strand labeled (asterisks, Fig. 2-3 and Fig. 2-4, lanes 3 to 6, respectively), suggesting that the

DNA might harbour some particular structure at these sites.

The translational setting of the nucleosomes was further investigated by restriction

enzyme analysis. Reconstituted nucleosomes were subjected to digestion with the restriction

enzymes PstI,AflIII, Hhaland Xhol, which cut 16, 31, 47/49 and 94 bps from the left end of the

30

2 Results

A/ Nucl. DNA A/ NucL DNA

Figure 2-3: ATDED-long nucleosome has a define rotational settingDNasel footprint ofATDED-long labeled on the top strand. The results of a short run (to the left) and a

long run (to the right) are shown. Digestion ofreconstituted nucleosomes (lanes 3 to 6) and naked DNA

(lanes 7 to 9) with DNasel. A/G (lane 1) and T (lane 2) are sequencing markers. Black dots mark

nucleosome-specific DNasel cutting sites; asterisks mark additional DNasel sensitive sites; numbers

show map units. The putative position of the histone octamer and the center of the nucleosome are

drawn schematically to the left of the gels.

DNA fragment, respectively (Fig. 2-1). On reconstituted fragments, cleavage by Xhol was

strongly inhibited (2% cut), in contrast to Afllll and Hhal, which restricted >80% of the DNA

and to Pstl, which restricted over 50% of the DNA (Fig. 2-5). Since digestion of reconstituted

nucleosomes by Pstl in other experiments restricted over 80% of the DNA (for example in

Fig. 2-36), we assume that the lower cutting efficiencies of Pstl compared to Afllll and Hhal

observed in Fig. 2-5 were due to some experimental parameter (amount or activity of the

enzyme) rather than to particular features of reconstituted nucleosomes.

31

2 Results

A/ Nucl. DNA A/ Nucl. DNA

"G T am ^ DNasel XC T ^ ^ DNasel

Figure 2-4: ATDED-long nucleosome has a define rotational settingDNasel footprint of ATDED-long labeled on the bottom strand. The results of a short run (to the left)

and a long run (to the right) are shown. Digestion of reconstituted nucleosomes (lanes 3 to 6) and naked

DNA (lanes 7 to 9) with DNasel. A/G (lane 1) and T (lane 2) are sequencing markers. Black dots mark

nucleosome-specific DNasel cutting sites; asterisks mark additional DNasel sensitive sites; numbers

show map units. The putative position of the histone octamer and the center of the nucleosome are

drawn schematically to the left ofthe gels.

Consistent with the DNasel analysis, the impeded cleavage observed at the Xhol site

indicates that the nucleosomes reconstituted on ATDED-long fragments were positioned toward

the right end of the DNA.

CPD formation in sequences containing T-tracts is modulated by folding of DNA in

nucleosomes (Schieferstein and Thoma, 1996). Since the ATDED-long sequence contains T-

tracts on both strands (Fig. 2-1), we further investigated reconstituted nucleosomes by UV

footprint analysis. DNA and nucleosomes were irradiated with UV at a dose of 750 J/m yielding

an average of 1.5 CPD per fragment. The DNA was isolated and damage formation was analysed

by digestion with T4-endoV and gel electrophoresis (Fig. 2-6). In naked DNA, the CPD

formation pattern was heterogen and reflected the structural properties of DNA, since the

32

2. Results

Top Strand Bottom Strand

DNA Nucl. DNA Nucl.

a- -«: a x

'HjHP jäfij^

%cut 96 99 99 99 51 84 79 2

%cut 94 100 62 85 92 2

100 100

Figure 2-5: Restriction enzymes accessibility assay

The results of the top (to the left) and bottom (to the right) strands are shown. Digestion of

reconstituted nucleosomes (Nucl. lanes) and naked DNA (DNA lanes) with the indicated

restriction enzymes. The cutting efficiencies are given in percent for each lane (bottom).

characteristic damage formation pattern of T-tracts (Lyamichev, 1991) was observed in clusters

2, 3, 4, 13 and 16 (top and bottom strands, lanes 2). In nucleosomal DNA, the CPD formation

was altered in the T-tracts of clusters 4, 13 and 16 compared with the CPDs formed in naked

DNA (top and bottom strands, lanes 4), indicating changes in the DNA structure upon folding

into nucleosomes. In the T-tracts of clusters 2 and 3, the CPD formation pattern did not change

upon reconstitution (top strand, lanes 2 and 4), consistent with their localisation outside of the

predicted nucleosome.

Taken together, we have a chromatin model substrate which consists of a positioned

nucleosome with a define rotational setting and about 60 bps of protruding naked "linker" DNA.

We used this model substrate to investigate CPD accessibility to the repair enzyme photolyase.

33

2 Results

Top Strand Bottom Strand

5'

DNAJNucl+ +

- +

DNA

+ + UV + +

- + T4EndoV -

Il m *»

IImi a

IT6

ZT5

~T9

12 3 4

3'

18

17

16

15

14

13

12 f

ggj|

T5

T8

12 3 4

Figure 2-6: UV footprint of ATDED-long nucleosomes

The results of the top (to the left) and the bottom (to the right) strands are shown.

ATDED-long DNA (lanes 1 and 2) or nucleosomes (lanes 3 and 4) were irradiated with

UV-light at a dose of 750 J/m2 and digested with T4-endoV (lanes 2 and 4). Nucleosome

and position of the damage clusters (to the left of the gels) and T-tracts of clusters 2, 3, 4,

13 and 16 (to the right of the gels) are indicated.

34

2. Results

2.1.2. Photoreactivation Is Modulated in ATDED-long Nucleosomes

Repair experiments were performed using irradiated nucleosomes. Irradiation of

nucleosomes with UV light at a dose of 750 J/m2 yielded an average of 1.5 CPDs per fragment.

Irradiated nucleosomes were incubated at 30°C with E. coli DNA photolyase in the presence of

photoreactivating light for up to 120 minutes. Band shift gels showed efficient incorporation of

labeled DNA into nucleosomes (>90%; Fig. 2-7, top and bottom strands, lanes 2). The

nucleosomal fraction remained unchanged over the course of UV irradiation and

photoreactivation (top and bottom strands, lanes 3 to 9). Hence, neither UV irradiation nor

photolyase disrupted nucleosomes.

Top Strand Bottom Strand

UV ^- + + + + + + +

<- + + + + + + +_

"

notoiyase p- -

5. 15. M< 45. m. 120' p~ ~

y is* »• «' w w

• -D

123456789 123456789

Figure 2-7: Native nucleoprotein gel analysis (containing 10% glycerol)Results of the top (to the left) and bottom (to the right) strands are shown. ATDED-long DNA

(lane 1) was reconstituted into nucleosomes (lane 2), irradiated with 750 J/m (lane 3) and

photoreactivated for 5, 15, 30, 45, 60 and 120 minutes (lanes 4 to 9, respectively). Bands

represent naked DNA (D) and nucleosomes (N).

We wanted to measure CPD repair in nucleosomal and naked DNA that had the same

damage distribution. Therefore, UV-irradiated nucleosomes and DNA isolated from these

irradiated nucleosomes were treated with photolyase. Following the reaction, DNA was purified

and the remaining CPDs detected by T4-endoV digestion and gel electrophoresis (Fig. 2-8 and

Fig. 2-9). Decreasing band intensities with increasing photoreactivation times reflect site-

specific repair by photolyase. Damages were quantified in CPD clusters numbered from 1 to 19

(Fig. 2-1). The percentage of repair is depicted either as function of the localisation of the

clusters on the DNA sequence (Fig. 2-10 B and Fig. 2-11 B) or as function of the

photoreactivation times (Fig. 2-10 C and Fig. 2-11 C).

Comparison of the initial damage and the repair rates of the individual clusters shows no

correlation between the amount of damages and the repair efficiency (Fig. 2-10, A and B and

Fig. 2-11, A and B).

35

2 Results

Nucl. DNA Nucl. DNA

5' 15' 30,45'60,120'5,10'15'30'45' PhOtOlyaSe "

y IS'30' 45'60'120' 5' 10' 15'30' 45'

- + + + + + + + + + + + ++ T4endoV - ++ + + + + + +++++ +

7 l&i.-,,* *^s S *** % *2* Jg| "^ "*• ;

8 tlflnislis9 *•*«*«,*•.»•«»-

2 4 6 8 10 12 141 3 5 7 9 H 13

1 2 3 4 5 6 7 8 9 10 11 12 13 14

Figure 2-8: Modulation of CPD repair in ATDED-long nucleosomes

Photoreactivation of ATDED-long labeled on the top strand. The results of a short run (to the left)and a long run (to the right) are shown. UV-irradiated nucleosomes (Nucl.) and DNA isolated from

these nucleosomes (DNA) were treated with photolyase for the indicated times (min.). Lane 2, initial

CPD distribution; lanes 4 to 9, photoreactivation in nucleosomes; lanes 10 to 13, photoreactivation in

naked DNA. Undigested DNA (lane 1) and overdigested DNA (lane 3) are controls. The position of

the histone octamer and the center of the nucleosome are drawn schematically to the left of the gels.The damage clusters that were quantified (see Fig. 2-10) are indicated (numbered from 1 to 9).

CPD repair in naked DNA was much faster than the corresponding repair rates in

nucleosomal DNA. In naked DNA, CPDs were repaired to completion within 45 minutes (Fig. 2-

8, lane 14; and Fig. 2-9, lane 15). In addition, repair of all damage clusters was rather

homogeneous (Fig. 2-10, B and C; and Fig. 2-11, B and C).

In reconstituted DNA, repair was fast outside of the nucleosome and slow in the

nucleosome. Repair of the clusters 1, 2, 3, 10 and 11, which are located outside of the

nucleosome, was almost complete after 30 minutes of photoreactivation (>75%), whereas the

clusters located in the nucleosome (clusters 5 to 9 and 13 to 19) remained poorly repaired

(<50%) even after 60 minutes of photoreactivation (Fig. 2-8, lanes 4 to 9 and Fig. 2-9, lanes 6 to

11; Fig. 2-10 B and Fig. 2-11 B). Similarly, the repair kinetics of the clusters located outside of

36

2 Results

19

1817

16

15

14

13

12

11

10

Nucl. DNA

+++++++++++++ UV

5' îs^o^s'amo'io'is^MS' "«OtOlyase+- + + + + + +++ + + + + T4endoV

Nucl. DNA

--++++++ + + + + + + +

5' 15' 30' 45' 60' 120' 10'15'30'45'

"+-++++++++ + + + +

HIi**s*M«»

*»*••«* —««•«*<*.—

Ï * * *

If!IfIff§5llfMllliMPII

119

18

17

I"lis

|u

I»112

H^ .^^ |gn -gàtt îgjgw jtafc| g^ .^y* ^at &jgk

*WÉ SÉÉ ^ÏÉÏ sSÉ: Mita i

AS»**«1* •

llWUli

2 4 6 8 10 12 141 3 5 7 9 11 13 15

2 4 6 8 10 12 1413 5 7 9 11 13 15

Figure 2-9: Modulation of CPD repair in ATDED-long nucleosomes

Photoreactivation ofATDED-long labeled on the bottom strand. The results of a short run (to the left)

and a long run (to the right) are shown. UV-irradiated nucleosomes (Nucl.) and DNA isolated from

these nucleosomes (DNA) were treated with photolyase for the indicated times (min.). Lane 4, initial

CPD distribution; lanes 6 to 11, photoreactivation in nucleosomes; lanes 12 to 15, photoreactivationin naked DNA. DNA isolated from non-irradiated nucleosomes (lanes 1 and 2), undigested DNA

(lane 3) and overdigested DNA (lane 5) are controls. The position of the histone octamer and the

center of the nucleosome are drawn schematically to the left of the gels. The damage clusters that

were quantified (see Fig. 2-11) are indicated (numbered from 10 to 19).

the predicted nucleosome (clusters 1, 2, 3, 10 and 11) were fast, while the clusters located in the

nucleosome (clusters 5 to 9 and 13 to 19) had slow repair kinetics (Fig. 2-10 C and Fig. 2-11 C).

Thus, the presence of a positioned nucleosome on the ATDED-long DNA had a localized

inhibitory effect on CPD repair by photolyase.

Clusters 4 and 12, which are located at the left edge of the nucleosome, showed an

intermediate repair efficiency, over 60% of the damages being repaired after 60 minutes

photoreactivation (Fig. 2-10 B and Fig. 2-11 B) and the repair kinetics being slightly slower than

37

2. Results

Initial CPDs

n

0.2

QOh

u

^ 0.10s-

n

n " nnfl n

B.

DNA

Nucl.

c.

100

I 50

0

100

.a

&50

0s-

1 2 34 56 7 89

15' 30' 45760'

•

I..IIIII),. - b.ll1 2 34 56789

Àdyade

DNA

1 2 34 56 7 89

dyade

1 2 34

Nucl.

56789

dyade

10 20 30

time [min]

40 20 40 60 80 100 120

time [min]

Figure 2-10:Quantification of CPD repair in the top strand of ATDED-long nucleosomes

A. Initial damage of clusters 1 to 9. B. Photoreactivation of clusters 1 to 9 was quantified in DNA

(DNA, empty bars) and nucleosomes (Nucl., black bars). Repair is shown as the fraction of CPD

removed by 15, 30 and 45 minutes of photoreactivation for naked DNA and by 15, 30 and 60 min.

of photoreactivation for nucleosomes. The position of the putative dyade is indicated. C.

Photoreactivation of clusters 1 to 9 was quantified in DNA (DNA) and nucleosomes (Nucl.).

Repair is plotted against photoreactivation time for each cluster.

38

2. Results

Initial CPDs

0.2

QCh

U0.1

m n ifl10

11

12141618

1315 17 19

B.

DNA

Nucl.

c.

100

2 50

100

50

-20

I

15' 30' 45760'

1 IllWirtln1-1 1- '-If-

Lfcji^ Jl^-1

(L10

11

1214 16 181315 17 19

dyade

DNA

10

il121416 181315 17 19

Adyade

1011

Nucl.

121416181315 17 19

Adyade

^-19

^18

•^-17+ 13

-^16

10 20 30

time [min]

40 20 40 60 80 100 120

time [min]

Figure 2-ll:Quantification of CPD repair in the bottom strand of ATDED-long nucleosomes

A. Initial damage of clusters 10 to 19. B. Photoreactivation of clusters 10 to 19 was quantified in

DNA (DNA, empty bars) and nucleosomes (Nucl., black bars). Repair is shown as the fraction of

CPD removed by 15, 30 and 45 minutes of photoreactivation for naked DNA and by 15, 30 and 60

min. of photoreactivation for nucleosomes. The position of the putative dyade is incicated. C.

Photoreactivation of clusters 10 to 19 was quantified in DNA (DNA) and nucleosomes (Nucl.).

Repair is plotted against photoreactivation time for each cluster.

39

2 Results

those of the 'linker' DNA clusters (Fig. 2-10 C and Fig. 2-11 C). Clusters 4 and 12 overlap with

the DNasel sensitive region observed around MU 1379 (Fig. 2-3 and Fig. 2-4), supporting the

idea that the DNA is generally accessible at this site.

Variation in repair efficiencies was observed within nucleosomal DNA. Repair has a

tendency to be more efficient with increasing distance from the putative dyade of the nucleosome

(Fig. 2-10 B and Fig. 2-11 B). Analysis of the repair kinetics of nucleosomal clusters reveals a

modulation of repair efficiencies in the predicted nucleosomes, some clusters (clusters 6, 7, 14,

15, 16) being resistant to repair and others (clusters 5, 8, 9, 13, 17, 18, 19) having slow repair

kinetics (Fig. 2-10 C and Fig. 2-11 C). No or very little increase of repair were observed for

photoreactivation times longer than 45 minutes.

2.1.3. The Rotational Setting of ATDED-long Nucleosomes Does not

Change upon UV-Irradiation and Photoreactivation

Previous studies showed that UV irradiation of pre-assembled nucleosomes does not alter

their structure (Liu et al., 2000; Schieferstein and Thoma, 1996). The nucleosomes analysed in

those studies were reconstituted on short DNA fragment. The ATDED-long sequence is 226 bps

in length and thus provides space for octamer movement along the DNA. Therefore, we

performed DNasel footprinting to investigate putative changes in the rotational setting of

ATDED-long nucleosomes after UV irradiation and during photoreactivation.

ATDED-long DNA was labeled on the top strand, reconstituted into nucleosomes,

irradiated with 750 J/m2 and photoreactivated for up to 180 minutes at 30°C. Native

nucleoprotein gel electrophoresis showed that the reconstitution efficiency was more than 90%

(Fig. 2-12 A, lane 2) and that the nucleosomes remained stable after UV-irradiation (lane 3) and

during photoreactivation for up to 120 minutes (lanes 4 to 6).

DNasel digestion ofATDED-long nucleosomes gave rise to the same nucleosomal cutting

pattern as described above (Fig. 2-12 B, lanes 3 and 4). No changes were observed in the

digestion pattern neither after UV irradiation (lanes 5 and 6) nor during up to 180 minutes

photoreactivation (lanes 7 to 14), indicating that the rotational setting of the nucleosomes

remained stable during the time course of the experiment.

2.1.4. Repair of Nucleosomal CPDs Remains Inhibited in the

Presence of Additional Photolyase

As described in Section 2.1.2., repair by photolyase is slow in the ATDED-long

nucleosome. Moreover, no or very little increase in repair were observed in samples incubated

for 45 minutes or longer (Fig. 2-10 C and Fig. 2-11 C). This results could be interpreted such that

40

2 Results

A.

uv «

Phr. g

B.

+ + + + +

30' 60' 120' ISO'

Nucl. DNA

+ + + + +

- 30' 60' 120' 180'

7GT

UV

Photolyase

DNasel

^^w^ ^^^ ~^d^ff^ ^P«|l^^

«ii

12 3 4 5 6 7

** jH' *R$F ^Wp tlP1 *"** '

fmfrJ§^ j^s» Mg ^§ jÉË :^M::^fe Mi Jl| jljbrt

IIîïîlîïlll?-"

^i^^ wi^ !•** ^^w !» ^^» ^^w ^^p -^n^-^m iipB ^* 1

: Jjjj| ^B igg

% -1362

-1374

1385

a *s& «* _^ 4* i«^ _ 1394

-«*•««««.»•»•*- -1404

«HlliflMlII- -1415

*I"

""«••_1426

e4**** •*•#•*• -1436

3m* S»

ifiSB S** t: - 1458

Üfjfe

« * • • .ino^iin «i» —1466

* jgji JE aft i „

f séa- als- *»s •s*»' ***

w ^^w ^^ ^^^^^^ ^^w ^—X't / O

1 2345 67 89 10 11 121314 15 16

Figure 2-12:DNaseI footprint after UV irradiation and during photoreactivationA. Nucleoprotein gel electrophoresis (containing 10% glycerol). ATDED-long DNA was labeled on the

top strand (lane 1), reconstituted into nucleosomes (lane 2), irradiated with UV light at a dose of 750 il

m (lane 3) and photoreactivated for the mdicated times (lanes 4 to 7). B. DNasel digestion of

nucleosomes (lanes 3 and 4), irradiated nucleosomes (lanes 5 and 6), nucleosomes photoreactivated for

the indicated times (lanes 7 to 14) and naked DNA (lanes 15 and 16). A/G (lane 1) and T (lane 2) are

sequencing marker. Black dots mark nucleosome-specific DNasel cutting sites; numbers show map

units. A schematic drawing of the nucleosome position is shown to the left of the gel.

the amount or the activity of the photolyase is a limiting factor for CPD repair, or that the enzyme

might be inactivated by long incubation times. To investigate this possibility, we performed the

following experiment. ATDED-long DNA was labeled on the bottom strand, reconstituted into

nucleosomes and irradiated with UV light at 750 J/m2 to yield 1.5 CPD per fragment on average.

41

2 Results

15' 30' 45' 60' 75' 90' 105' 120'

+ + + +

+

+

+

+

+

The irradiated nucleosomes were split in two samples. Both samples were photoreactivated as

described previously. However, fresh photolyase was added to one sample after 45 and 90

minutes and the reaction incubated for up to 120 minutes.

UV -+++++++++

Photolyase ^

JS«jl JÈ&L Jä||| |jM| gj^^ ^%fk :^H||. jgg|g; $^a!ÊL.Ê^^k__ îyr

^Hipp ^H^ !^^^P? SHIIP ^piw ^^^^ ^HIP? ^Q^^ ^^^gß~ f^^^w A~

:«||||: ««til»- •*** *< » -s*~ w m, *• & «##— D

123456789 10 11

Figure 2-13:Native nucleoprotein gel analysis (containing 10% glycerol)

ATDED-long DNA was labeled on the bottom strand (lane 1), reconstituted into

nucleosomes (lane 2), irradiated with 750 J/m2 (lane 3) and photoreactivated for 15, 30, 45,

60, 90, 105 and 120 minutes (lanes 4 to 10, respectively). Fresh photolyase was added after

45 and 90 minutes. Bands represent naked DNA (D) and nucleosomes (N).

Native nucleoprotein gel electrophoresis showed that the nucleosomes remained stable

over the course of the experiment (Fig. 2-13, lanes 3 to 11). CPD repair was analysed (Fig. 2-14)

and quantified (Fig. 2-15).

Direct comparison (panel A, empty vs. black bars) showed that both nucleosomal samples

were repaired at comparable rates prior to the addition of fresh photolyase (45 minutes). After

addition of fresh photolyase (60 minutes), repair of most ofthe damage clusters remained similar

in both samples, whereas the repair of clusters 13, 14 and 15 was increased in the sample

containing additional photolyase. The repair curves (panel B) confirm that repair of clusters 13,

14 and 15 was enhanced after addition of fresh photolyase, since the slope of their curves

increased after the first addition of photolyase (45 minutes). After 120 minutes of

photoreactivation and two additions of photolyase, cluster 19 was efficiently repaired (>75%),

while repair of clusters 13 to 18 remained slow and inefficient (<50%). The resulting modulation

of repair can be summarized as follows: the clusters located outside of the predicted nucleosome

(clusters 10 and 11) as well as two most distal clusters in the nucleosome (clusters 12 and 19)

were efficiently repaired, while repair of the nucleosomal clusters 13 to 17 was slow and

inefficient. Thus, the addition of fresh photolyase is not sufficient to achieve complete repair of

nucleosomal CPDs.

42

2 Results

Nucl. Nucl. DNA

-+++++++++ +++ + +++++++

19

1817

16

15

14

13

12

111

110

UV

Photolyase

Nucl. Nucl. DNA

5' 15'30 45 60* 15' 30'45'60* 75* 90*105*120*5*10 15 »' DUAeftUrncA 5' 15'30 45'60'15'30'45* 60'75' »0* 105*120'5* 10' 1530'

++++++rnotoiyase

++++++

+++ +++

+ -+++++++ ++++ + +++++ + + T4endoV - + -+++++++++++++++++ ++

liliirtits:--*-!!*

|£ m éê mm

2 4 6 8 10 12 14 16 18 20 221 3 5 7 9 11 13 15 17 19 21

119

18

I 17

I"lis

I ,4

I"12

filial»!»* -». Mm-*

M- <»* W* #. <*• «z ®» m» «& *s &> m« <** i&« & m* f

wViIWmP^fIPWWm

ff§•* §«*•#-»ft**»"*

IHiHIHHiWIi-

»ffc«_ » 4,- ,

2 4 6 8 10 12 14 16 18 20 221 3 5 7 9 11 13 15 17 19 21

Figure 2-14:Photoreactivation of ATDED-long nucleosomes

Photoreactivation ofATDED-long labeled on the bottom strand. The results of a short run (to the left)

and a long run (to the right) are shown. UV-irradiated nucleosomes (Nucl.) and DNA isolated from

these nucleosomes (DNA) were treated with photolyase for the indicated times (min.). Where indicated,

fresh photolyase was added to the sample. Lane 4, initial CPD distribution; lanes 6 to 10,

photoreactivation in nucleosomes; lanes 11 to 17, photoreactivation in nucleosomes with additional

photolyase; lanes 18 to 21, photoreactivation in naked DNA. DNA isolated from non-irradiated

nucleosomes (lanes 1 and 2), undigested DNA (lane 3) and overdigested DNA (lane 5) are controls. The

position ofthe histone octamer and the center of the nucleosome are drawn schematically to the left of

the gels. The damage clusters that were quantified (see Fig. 2-15) are indicated (numbered from 10 to

19).

43

2 Results

A.100

a so

0

-10

45' 60'

lU Jii10

il121416181315 17 19

10

11121416181315 17 19

B. + phr + phr

TIA-x-12

-«-19

^18"^13

^15-»-17

-e-14

h^-16

0 20 40 60 80

time [mm]

100 120 0 20 40 60 80

time [mm]

100 120

Figure 2-15:Quantification of CPD repair in ATDED-long nucleosomes

Photoreactivation of clusters 10 to 19 was quantified in nucleosomes. A. Repair is shown as the

fraction of CPD removed after 45 and 60 minutes photoreactivation in the control samples (white

bars) and in the samples in which additional photolyase was added (black bars). B. Repair is plotted

against photoreactivation time for each cluster.

44

2. Results

2.2. ySWI/SNF Remodeling ofATDED-long Nucleosomes

Recent studies indicate that ATP-dependent chromatin remodeling facilitate transcription

and others processes having chromatin as substrate through local alteration of nucleosomes

(reviewed in (Fyodorov and Kadonaga, 2001; Narlikar et al., 2002)). Here, we used the ySWI/

SNF complex and our model substrate to investigate the effect of nucleosome remodeling on

DNA damage formation and repair in nucleosomes.

2.2.1. Remodeling by ySWI/SNF Alters the Structure of Nucleosomes

To test the activity of the ySWI/SNF complex in our model substrate, DNasel and UV

footprint analyses were performed. ATDED-long nucleosomes, end-labeled at the bottom strand,

were incubated with ySWI/SNF either in the absence or in the presence ofATP prior to DNasel

or UV footprinting.

In the DNasel footprint analysis, a molar ratio of one ySWI/SNF to one nucleosome was

used. Native nucleoprotein gels showed that the labeled ATDED-long nucleosomes were

completely bound by ySWI/SNF to form a complex, which was too big to enter in the gels and

therefore remained stuck in the wells (Fig. 2-16, lanes 3 and 4).

Figure 2-16:Native nucleoprotein gel electrophoresis

(containing 10% glycerol)

ATDED-long DNA was end-labeled on the bottom strand

(lane 1), reconstituted into nucleosomes (lane 2) and

incubated for 30' at 30°C with ySWI/SNF (lanes 3 and 4)

and ImM ATP (lane 4). The molecular ratio of SWI/SNF

to nucleosomes was of 1. Bands represent naked DNA

(D), nucleosomes (N) and the wells (W).

DNasel footprinting showed that the rotational setting of the nucleosome got lost after

incubation with ySWI/SNF and ATP (Fig. 2-17, lanes 9 and 10). The resulting cutting patterns

looked more similar to naked (lanes 2 to 4) than to nucleosomal DNA (lanes 5 and 6). The

nucleosomal pattern was not altered in the absence of ATP (lanes 7 and 8), except around MU

1377, where the cutting pattern was more equal than in reconstituted nucleosomes, suggesting

+ - - - DNA

+ + + Nucl.

- - + + SWI/SNF

- - - + ATP

we

-N

-D

45

2 Results

Figure 2-17:Altered nucleosome structure upon remodeling by ySWI/SNFDNasel footprinting of the remodeled nucleosomes described in Fig. 2-16. The results of a short run

(to the left) and a long run (to the right) are shown Digestion of naked DNA (lanes 2 to 4),

nucleosomes (lanes 5 and 6), nucleosomes incubated with ySWI/SNF in the absence (lanes 7 and 8)

or the presence of 1 mM ATP (lanes 9 and 10) with DNasel. A/G (lane 1) is a sequencing marker.

Black dots mark nucleosome-specific DNasel cutting sites; asterisks mark additional DNasel

sensitive sites; numbers show map units. The position of the histone octamer and the center of the

nucleosome are drawn schematically to the left of the gels.

that ySWI/SNF binding might protect the DNA from digestion at these sites. This region

correspond to the DNasel sensitive site of the ATDED-long nucleosome (see Section 2.1.1.),

which is efficiently repaired by photolyase (see Section 2.1.2.).

46

2. Results

Thus ySWI/SNF seems to behave on ATDED-long nucleosomes as previously described

for other substrates with respect to DNasel accessibility (Cote et al., 1998; Cote et al., 1994;

Owen-Hughes et al., 1996).

Since reconstitution of the ATDED-long sequence in nucleosomes changes the CPD

formation pattern (Fig. 2-6), we investigated the structural changes induced by incubation with

ySWI/SNF on ATDED-long nucleosomes by UV-footprinting.

Due to the limited amount ofySWI/SNF available, the molar ratio had to be reduced to 0.8

ySWI/SNF to one nucleosome in the UV footprint experiment compared to the DNasel

experiment, where an equimolar ratio was used. Native nucleoprotein gel electrophoresis showed

that the labeled ATDED-long nucleosomes were bound by ySWI/SNF (Fig. 2-18, lanes 3 and 4),

generating a smear along the lanes and toward the wells.

+ - +

+ + + + + +

+ + - - + +

- + - - - +

- - + + + +

DNA

^. Figure 2-18:Native nucleoprotein gel of

nucleosomes remodeled by ySWI/SNFSWI/SNF (containing 10% glycerol)ATP

_

ATDED-long DNA was end-labeled on

plasmid DNA the bottom strand (lane 1), reconstituted

#» -W

-N'

-N

into nucleosomes (lane 2) and incubated

for 30' at 30°C with ySWI/SNF (lanes 3

and 4) and ImM ATP (lane 4). The

molecular ratio of ySWI/SNF to

nucleosomes was of 0.8. Lanes 5 to 8

show the samples described above after

competition of ySWI/SNF. An aliquot of

the samples loaded in lanes 1 to 4 were

incubated with plasmid DNA in excess for

30' at 30°C. Bands represent naked DNA

(D), nucleosomes (N), an unidentified new

band (N') and the wells (W).

To test the state of the nucleosomes after incubation with ySWI/SNF and prior to UV-

irradiation, aliquots were incubated with an excess of plasmid DNA to compete ySWI/SNF away

from the nucleosomes and analysed by native nucleoprotein gel electrophoresis (Fig. 2-18, lanes

5 to 8). No significant changes in the relative proportions of recovered nucleosomes vs. naked

DNA were observed, in agreement with previous work, where no or very little increase in naked

DNA were observed in similar competition experiments (Cote et al., 1998; Guyon et al., 1999;

Owen-Hughes et al., 1996; Sengupta et al., 2001). However, in addition to the naked (D) and

nucleosomal (N) bands, a novel, slower migrating band appeared independently of the presence

47

2 Results

ofATP (N', lanes 7 and 8). Since the nucleoprotein gel contained 10% glycerol, it is unlikely that

the novel band consists of new positions of the histone octamer on the DNA fragment (Pennings

et al., 1992). Furthermore, nucleosomes positioned centrally on the fragment would not be

expected to migrate so much slower than end-positioned nucleosomes (see Section 2.3.). The

slower migrating band resembles the novel species described in previous studies with yRSC

(Lorch et al., 1998) and with hSWI/SNF (Schnitzler et al., 1998), which were identified as

dinucleosomes. The appearance of dinucleosomes was also observed with ySWI/SNF, although

not consistently (Sengupta et al., 2001). The appearance of slower migrating, dinucleosome-like,

bands might depend on the experimental conditions (type of nucleoprotein gel, nucleosome

concentration, length of the DNA, kind of competitor).

For UV photofootprinting, the samples were irradiated with UV at a dose of 500 J/m

yielding an average of 1 CPD per fragment. The DNA was isolated and damage formation was

analysed by digestion with T4-endoV and gel electrophoresis (Fig. 2-19).

In nucleosomal DNA, the CPD formation was altered in the T-tracts of clusters 13 and 16

compared to the CPDs formed in naked DNA (Fig. 2-19, lanes 4 and 5), indicating changes in the

DNA structure upon folding into nucleosomes, as observed above (Fig. 2-6). After incubation

with ySWI/SNF and ATP, the CPD formation pattern resembled that of naked DNA. The

structural alteration was due to the remodeling activity of ySWI/SNF since addition of the

complex in the absence of ATP did not alter the nucleosomal CPD pattern (lane 6). Our results

indicate a change in the structure of nucleosomal DNA upon remodeling by ySWI/SNF.

2.2.2. Enhanced CPD Repair Following ySWI/SNF Nucleosome

Remodeling

Since the positioned ATDED-long nucleosome had a strong inhibitory effect on CPD

repair by photolyase and that the nucleosome structure was altered by ySWI/SNF, we wanted to

analyse the effect of remodeling on photoreactivation. Nucleosomes were irradiated with UV

light at a dose of 500 J/m2, yielding one CPD/fragment on average. The irradiated nucleosomes

were incubated with ySWI/SNF either in the absence or in the presence ofATP and exposed to

E.coli DNA photolyase in the presence of photoreactivating light for up to one hour. The molar

ratio was 0,6 ySWI/SNF per nucleosome.

Nucleoprotein gel electrophoresis (Fig. 2-20 A) showed that reconstitution was efficient

(>85%, lane 2) and that the nucleosomes were stable over the course of the experiment (lanes 3

to 7). Addition of ySWI/SNF resulted in a nearly complete shift of the nucleosomes in high

48

2 Results

+.+... DNA

- + - + + + Nucl.

....++ SWI/SNF

A, + ATP'G . . + + + + T4EndoV

Figure 2-19:UV-footprint of nucleosomes

remodeled by ySWI/SNFNaked DNA (lane 4), nucleosomes (lane 5),

nucleosomes incubated with ySWI/SNF in

the absence (lane 6) or the presence of 1 mM

ATP (lane 7) were irradiated with 500 J/m2.

The molecular ratio of ySWI/SNF to

nucleosomes was of 0.8. A/G (lane 1) is a

sequencing marker. Undigested DNA (lane

2) and nucleosomes (lane 3) are controls.

Nucleosome and position of the damageclusters (to the left) and T-tracts of clusters

13 and 16 (to the right) are indicated.

molecular weight complexes, that appeared as a smear or remained stuck in the wells (lanes 8 to

15), indicating that ySWI/SNF binds to damaged nucleosomes. Similar binding occurred in the

absence and in the presence ofATP (compare lanes 8 to 11 with lanes 12 to 15).

To analyse the state of the remodeled nucleosomes over the course of the experiment,

aliquots were taken and incubated with an excess of plasmid DNA to compete for ySWI/SNF

(Fig. 2-20 B). In the absence of ySWI/SNF, the nucleosomes remained unaffected (lanes 4 to 7).

After incubation with ySWI/SNF in the absence ofATP, addition of plasmid DNA resulted in the

recovery ofthe nucleosomal band (lanes 8 to 11), indicating that the addition of competitor DNA

was sufficient to compete ySWI/SNF away from the labeled nucleosomes. In the presence of

ATP, competition with plasmid DNA was not complete (lanes 12 to 15). About 17-25% of the

49

2. Results

> _ +

p

< "3 "3- -

z a a ^^rt

a Z Z is' so¬ ar 15' 30' 60'

+

+

15' 30' 60'

ySWI/SNFATP

Photolyase••Mfc* »*•«•** ** -W

MMHIM-N

-D

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15

B. - + + ySWI/SNF

^ - - + ATP

DNA Nucl. Nucl. 15' 30' 60'"

15' 30' 60'-

is- 30' Z Photolyase

+ + + + + + + + + + + + Plasmid DNA

r * •>•"• m> • -w

mm Mm* * —N

-D

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15

Well [%] 1 1 3

Nucl. [%] 86 92 95-96

DNA [%] 13 6 1-3

4-6

92-95

1

17-25

66-77

6-9

Figure 2-20:Remodeling ofATDED-long nucleosomes by ySWI/SNFA. Nucleoprotein gel analysis. ATDED-long DNA was end-labeled on the bottom strand (lane

1), reconstituted into nucleosomes (lane 2) and irradiated with 500 J/m2 (lane 3). Irradiated

nucleosomes were incubated alone (lane 4), with ySWI/SNF (lanes 8 and 12) and 0.5 mMATP

(lane 12) for 30' at 30°C and photoreactivated for the indicated times (lanes 5 to 7, 9 to 11 and

13 to 15, respectively). The molecular ratio of ySWI/SNF to nucleosomes was of 0,6. Bands

represent naked DNA (D), nucleosomes (N) and the wells (W). B. Nucleoprotein gel analysisafter competition ofySWI/SNF. The samples described in panel A were incubated with plasmidDNA in excess for 45' at room temperamre. Quantification of the signal in the well (W), the

nucleosomal (N) and the naked DNA (D) bands is indicated for each lane (bottom).

50

2. Results

labeled DNA remained stuck in the wells, 66-77% migrated as nucleosomes and 6-9% as naked

DNA. These results indicate that most of the DNA remained folded into nucleosomes after

remodeling by ySWI/SNF and over the course of photoreactivation.

To allow direct comparison of CPD repair, DNA isolated from irradiated nucleosomes,

irradiated nucleosomes and irradiated nucleosomes incubated with ySWI/SNF were treated with

photolyase for various times. The remaining CPDs were analysed (Fig. 2-21) and quantified in

CPD clusters numbered from 10 to 19 (Fig. 2-1). The percentage of repair is depicted either as

function of the localisation of the clusters on the DNA sequence (Fig. 2-22) or as function of the

photoreactivation times (Fig. 2-23).

Repair of naked DNA was homogeneous and complete after 30 minutes photoreactivation

(Fig. 2-21, lanes 15 to 17; Fig. 2-22 and Fig. 2-23) whereas repair of nucleosomes was

modulated (Fig. 2-21, lanes 6 to 8; Fig. 2-22 and Fig. 2-23), as shown above (Fig. 2-11).

Incubation with ySWI/SNF in the absence of ATP did not alter the overall repair pattern

(Fig. 2-21, lanes 9 to 11; Fig. 2-22 and Fig. 2-23). Repair remained fast in clusters 10 and 11 and

inefficient in clusters 13 to 19. However, the repair rates of single clusters in the positioned

nucleosome were changed upon ySWI/SNF binding. Repair was reduced in some clusters (12, 13

and 19) and increased in others (16 to 18; Fig. 2-22). The resulting repair kinetics of the clusters

located in the nucleosome appeared less heterogeneous than in the absence ofySWI/SNF (Fig. 2-

23). These results indicate that the accessibility of some nucleosomal CPDs was slightly

modulated upon ySWI/SNF binding.

In the presence of ySWI/SNF and ATP, substantial changes in repair were observed

(Fig. 2-21, lanes 12 to 14; Fig. 2-22 and Fig. 2-23). Efficient repair was observed in clusters 10

to 18 (>50% after 60' photoreactivation; Fig. 2-22), while cluster 19 remained poorly repaired

(30% after 60' photoreactivation). After incubation with ySWI/SNF and ATP, repair of clusters

10 and 11 was reduced while repair of clusters 12 to 18 increased, resulting in a more

homogeneous repair than in nucleosomes. Correspondingly, the repair kinetics appeared rather

homogeneous in the repair curves (Fig. 2-23).

Our results demonstrate that incubation ofATDED-long nucleosomes with ySWI/SNF and

ATP had a profound effect on the accessibility of CPDs to photolyase. By increasing the

accessibility of nucleosomal CPDs, chromatin remodeling could provide a general mechanism to

bypass the inhibitory effect of nucleosomes on DNA repair. This finding led us to ask whether

other ATP-dependent remodeling complexes also alter CPD repair on nucleosomes.

51

2 Results

+ + +

+ - + +

19

1817

16

15

14

13

12

+

Nucl.

+ +

+

15' 30' 60' 15' 30' 60' 15' 30' 60'

+ + + + +++ + +

+ UV - - + + +

DNA

- SWI/SNF

ATP

— Phr

15' 30' 60'x '"

+ ± + T4endoV - + - + +

+ +

Nucl. DNA

+ + -

+

15' 30' 60' 15' 30' 60' 15 30' 60' 15' 30' 60'

+ + + + + +++ + + + +

•llïhlÉÉÉ IM àtt ^msl& ^^ ütt ai Mi *

•H Aü **t mu tet^ jgg iHü& *

X X mm ««s*

123456789 10 11 1213 14151617

|19

18

17

I"|,

I»

I»112

.m ^ m ÉIIÊ* <t BS ^

m, g» «•«*<•**•»•*'*

» d« f- t

"

R ff*

- -« »» « "* lif W*

mmmMàa "AMI

sffP»W"HP*"

iiiiiÈl p*.

1 2 3 4 5 6 7 8 9 10 11 12 13 14 1516 17

Figure 2-21:Photoreactivation of nucleosomes remodeled by ySWI/SNFPhotoreactivation of ATDED-long labeled on the bottom strand. The results of a short (to the left) and a

long run (to the right) are shown. DNA isolated from UV-irradiated nucleosomes (lanes 15 to 17), UV-

irradiated nucleosomes (lanes 6 to 8) and UV-irradiated nucleosomes incubated with ySWI/SNF (lanes 9

to 11) and 0.5 mM ATP (lanes 12 to 14) were treated with photolyase for the indicated times (min.). Lane

4, initial CPD distribution. DNA isolated from non-irradiated nucleosomes (lanes 1 and 2), undigestedDNA (lane 3) and overdigested DNA (lane 5) are controls. The position of the histone octamer and the

center of the nucleosome are drawn schematically to the left of the gels. The damage clusters that were

quantified (see Fig. 2-22 and Fig. 2-23) are indicated (numbered from 10 to 19).

52

2. Results

DNA

100

•a

S 50

S?

Nucl.ao,!»

0

100

50

Nucl.

SWI/SNF

o-10

100

50

0-10

100

Nucl. |

SWI/SNF §* so

ATP^

15' 30* 60'

J

njnurfl iWIlll nJlJ

^lSl Jj obi

121416181315 17 19

10 121416 1811 1315 17 19

10 1214161811 1315 17 19

Figure 2-22:Quantification of repair in remodeled nucleosomes

Quantification of CPD repair by photolyase of Fig. 2-21. Photoreactivation of clusters 10 to 19 was

quantified in naked DNA (grey bars), nucleosomes (empty bars) and in nucleosomes incubated with

ySWI/SNF (dash bars) and ATP (black bars). Repair is shown as the fraction of CPD removed by 15,

30 and 60 min. of photoreactivation.

53

2. Results

DNA Nucl.

0 10 20 30 40

time [min]

50 0 10 20 30 40

time [min]

-^-19^-18-«-17H3-16-e-15-B-14-^13-*-12^-11-«-10

Nucl. + SWI/SNF Nucl. + SWI/SNF + ATP

0 10 20 30 40

time fmin]

50 60 10 20 30 40 50 60

time [mini

Figure 2-23:Quantification of repair in remodeled nucleosomes

Quantification ofCPD repair by photolyase of Fig. 2-21. Photoreactivation of clusters 10 to 19

was quantified in naked DNA (DNA), nucleosomes (Nucl.) and in nucleosomes incubated with

ySWI/SNF (Nucl.+SWI/SNF) and ATP (Nucl.+SWI/SNF+ATP). Repair of each cluster is

plotted as function of incubation time.

54

2. Results

2.3. yISW2 Remodeling ofATDED-long Nucleosomes

The yISW2 complex has been shown to burn ATP in the presence of nucleosomes and to

have nucleosome spacing activities in vitro as well as in vivo (Kent et al., 2001; Tsukiyama et al.,

1999). Here, we used the yISW2 complex and our model substrate to investigate the effect of

nucleosome remodeling on CPD recognition and repair in nucleosomes.

2.3.1. yISW2 Alters CPD Repair

To test the effect of yISW2 on CPD accessibility and repair, the following

photoreactivation experiments were performed on ATDED-long nucleosomes. Nucleosomes

labeled on either strand were irradiated with UV light at a dose of 500 J/m2, incubated with

yISW2 either in the absence or in the presence ofATP and exposed to E.coli DNA photolyase in

the presence of photoreactivating light for up to one hour. The molar ratio was 0.7 yISW2 per

nucleosome and the nucleosome concentration was 20 ng/ul.

Nucleoprotein gel electrophoresis (Fig. 2-24) showed that the efficiency of reconstitution

was >80%, that the nucleosomes migrated prominently as one band (Nl) and were stable over

the course of the experiment (lanes 2 to 14). In the presence of yISW2, a minor fraction of the

labeled nucleosomes was shifted and appeared as a smear or remained stuck in the wells (lanes 7

to 14), indicating the that the amount of complex used was not sufficient to firmly bind all

nucleosomes or that binding occurred transiently. Incubation with yISW2 in the absence ofATP

did not change the migration of the nucleosomes over the course of the experiment (lanes 7 to

10).

In the presence ofATP, the mobility of the nucleosomes was reduced (N2; lanes 11 to 14)

compared to the mobility of nucleosomes incubated in the absence of ATP (lanes 7 to 10) and

yISW2 (lanes 2 to 6). Although the amount of yISW2 used shifted only a minor fraction of the

nucleosomes, it was sufficient to reduce the migration of the majority of the labeled nucleosomes

in the presence of ATP. The nucleosomes kept their reduced mobility over the course of

photoreactivation.

Since nucleosomes located at a fragment end migrate faster than those that are positioned

more centrally on the fragment in non-denaturing gels (Linxweiler and Horz, 1984; Pennings et

al., 1991), it is possible that the reduced mobility observed upon incubation with yISW2 and ATP

was due to a shift of the nucleosomes to more central positions.

55

2 Results

>

< 73£ s

Top Strand

+

15' 30' 60' 15' 30' 60'

+ ISW2

+ ATP

15- 30. 7 Photolyase

IPJPjPf ^PJPP ^^Sp^^S|p "^p^B8 ^^^w ^^^P1 ^^^P Jl| J.

-D

1 2 3 4 5 6 7 8 9 10 11 12 13 14

>

< -àfc 3

Bottom Strand

+

15' 30' 60' I~

15' 30' 60'

+ ISW2

+ ATP

15. 30' T Photolyase

j^^^^lj^;^^(|^|^^J^Éil^lSiP' —N2

-D

12 3 4 5 6 7 8 9 10 11 12 13 14

Figure 2-24:Native nucleoprotein gel analysisThe results of the top (upper panel) and the bottom (lower panel) strands are shown. ATDED-

long DNA (lane 1) was reconstituted into nucleosomes and irradiated with 500 J/m2 (lane 2).

Irradiated nucleosomes were incubated alone (lane 3), with yISW2 (lanes 7) and 0.5 mMATP

(lane 11) for 30' at 30°C and photoreactivated for the indicated times (lanes 4 to 6, 8 to 10 and

12 to 14, respectively). The molecular ratio was of 0.7 yISW2 per nucleosomes and the

nucleosome concentration was 20 ng/ul in all reactions. Bands represent naked DNA (D),nucleosomes (Nl) and nucleosomes with reduced mobility (N2).

To allow direct comparison ofCPD repair, irradiated nucleosomes, irradiated nucleosomes

incubated with yISW2 and DNA isolated from irradiated nucleosomes were treated with

photolyase for various times. The remaining CPDs were analysed (Fig. 2-25 and Fig. 2-26) and

56

2 Results

quantified in CPD clusters numbered from 1 to 19 (Fig. 2-1). The percentage of repair is depicted

either as function of the localisation of the clusters on the DNA sequence (Fig. 2-27 and Fig. 2-

29) or as function of the photoreactivation times (Fig. 2-28 and Fig. 2-30).

+ + +

- + - + +

+

Nucl.

+ +

+

15' 30' 60' 15' 30' 60' 15' 30' 60'

+ +++ + + + + +

+ + +

15' 30'60'

+ + + T4endoV - + - + +

+ +

Nucl. DNA

+ + -

+

15' 30' 60' 15' 30' 60' 15' 30'60' 15' 30' 60'

+ ++++++• + + + + +

i

I

:*Ul**)tS! ""'

12 3 4 5 6 7 8 9 1011121314 151617

123456789 1011121314 151617

Figure 2-25:Photoreactivation of nucleosomes remodeled by yISW2Photoreactivation in naked DNA, nucleosomes and remodeled nucleosomes labeled on the top strand.

The results of a short (to the right) and a long (to the left) run are shown. DNA isolated from UV-

irradiated nucleosomes (lanes 15 to 17), UV-irradiated nucleosomes (lanes 6 to 8) and UV-irradiated

nucleosomes incubated with yISW2 (lanes 9 to 11) and 0.5 mM ATP (lanes 12 to 14) were treated with

photolyase for the indicated times (min.). The molecular ratio was of 0.7 yISW2 per nucleosomes and

the nucleosome concentration was 20 ng/ul in all reactions. Lane 4, initial CPD distribution. DNA

isolated from non-irradiated nucleosomes (lanes 1 and 2), undigested DNA (lane 3) and overdigestedDNA (lane 5) are controls. The position of the histone octamer and the center of the nucleosome are

drawn schematically to the left of the gels. The damage clusters that were quantified (see Fig. 2-27 and

Fig. 2-28) are indicated (numbered from 1 to 9).

57

2 Results

+ + +

- + - + +

+

Nucl.

+ +

+

15' 30' 60' 15' 30' 60' 15' 30' 60'

+ + ++ + + ++ +

+

DNA

UV - - + + +

ISW2

ATP

^ Phr

15' 30' 60'A "*

+ + + T4endoV - + - + +

+

Nucl.

+ +

+

15' 30' 60' 15' 30'60' 15' 30' 60'

+ ++ + ++++ +

+

DNA

15' 30' 60'

+ + +

123456789 10111213141516 17

19

14

13

112

18 »MM»»«»»«-:::

|17n is* ä ^ it§ ^

1 sig - a ssi <*i «a

1" Kllilllgig

|.5 sitiplaaia MB

aaaataa-aaf

2 3 4 5 6 7 8 9 1011121314151617

Figure 2-26:Photoreactivation of nucleosomes remodeled by yISW2Photoreactivation in naked DNA, nucleosomes and remodeled nucleosomes labeled on the bottom

strand. The results of a short (to the right) and a long (to the left) run are shown. DNA isolated from UV-

irradiated nucleosomes (lanes 15 to 17), UV-irradiated nucleosomes (lanes 6 to 8) and UV-irradiated

nucleosomes incubated with yISW2 (lanes 9 to 11) and 0.5 mM ATP (lanes 12 to 14) were treated with

photolyase for the indicated times (min.). The molecular ratio was of 0.7 yISW2 per nucleosomes and

the nucleosome concentration was 20 ng/ul in all reactions. Lane 4, initial CPD distribution. DNA

isolated from non-irradiated nucleosomes (lanes 1 and 2), undigested DNA (lane 3) and overdigestedDNA (lane 5) are controls. The position of the histone octamer and the center of the nucleosome are

drawn schematically to the left of the gels. The damage clusters that were quantified (see Fig. 2-29 and

Fig. 2-30) are indicated (numbered from 10 to 19).

Repair of naked DNA was homogeneous and complete after 30 minutes photoreactivation

(Fig. 2-25 and Fig. 2-26, lane 16; Fig. 2-27 and Fig. 2-29) whereas repair of ATDED-long

nucleosomes was modulated (Fig. 2-25 and Fig. 2-26, lanes 6 to 8; Fig. 2-27 and Fig. 2-29) as

shown above (Fig. 2-10 and Fig. 2-11). Incubation with yISW2 in the absence of ATP did not

58

2. Results

alter the overall repair pattern (Fig. 2-25 and Fig. 2-26, lanes 9 to 11 ; Fig. 2-27 and Fig. 2-29).

An increase of nucleosomal repair was observed in clusters 6 to 8 of nucleosomes labeled on the

top strand (Fig. 2-27), resulting in more homogeneous repair (Fig. 2-28). However, no

significant increase was observed in nucleosomes labeled on the bottom strand (Fig. 2-29), in

which the repair kinetics of the individual clusters were very similar to those observed in the

absence of yISW2 (Fig. 2-30).

In the presence of yISW2 and ATP, repair of individual clusters were altered with either

strand labeled (Fig. 2-25 and Fig. 2-26, lanes 12 to 14; Fig. 2-27, Fig. 2-28, Fig. 2-29 and Fig. 2-

30). On the top strand, clusters 8 and 9 became more (>60% repair after 15') and clusters 2, 3,4

and 5 less accessible to photolyase upon incubation with yISW2 and ATP (Fig. 2-27). Repair was

efficient in clusters 1, 2, 3, 8 and 9 (>50% repair after 30') while clusters 4, 5, 6 and 7 remained

poorly repaired (<40% repair after 60'), resulting in a modulated repair pattern in which the

repair of clusters located centrally on the DNA fragment were inhibited. Correspondingly, the

repair kinetics of clusters 1, 2, 3, 8 and 9 were rather fast, while clusters 4, 5, 6 and 7 showed

slow repair kinetics (Fig. 2-28).

On the bottom strand, clusters 14 to 19 became more (>30% repair after 60') and clusters

12 and 13 less (<30% repair after 60') accessible to photolyase upon incubation with yISW2 and

ATP (Fig. 2-29). Repair of clusters 10, 11, 18 and 19 was fast (>70% after 60'), while repair of

clusters 12 to 17 was inefficient (<40% after 60'), resulting in a modulated repair pattern, with

the inhibited region located toward the center of the fragment. In contrast to the results obtained

with ySWI/SNF, the repair kinetics of nucleosomes incubated with yISW2 and ATP were as

heterogeneous as in ATDED-long nucleosomes, although the repair rate of individual clusters

were modified (Fig. 2-30).

Thus, repair analysis of both strands are consistent with a movement of the nucleosomes

toward more central positions upon remodeling by yISW2.

The repair pattern obtained on nucleosomes remodeled by yISW2 was different from the

repair pattern observed after remodeling by ySWI/SNF (compare Fig. 2-29 to Fig. 2-22). While

the ySWI/SNF complex generally increased the accessibility of nucleosomal CPDs to

photolyase, the yISW2 complex apparently mobilized the nucleosomes to more central positions.

Our results show that two ATP-dependent remodeling complexes, ySWI/SNF and yISW2,

remodel UV irradiated nucleosomes and that remodeling by either of the complexes can facilitate

CPD repair in nucleosomal DNA.

59

2 Results

100

DNA 'S 50

Nucl.

Nucl.

ISW2

0

100

I»^o^

0

100

t50

0s

0

100

Nucl.u

ISW2 'g.50ATP |

0s-

15* 30' 60'

'

nn nr d] i t -

I"l

ninnfl LiJI: Llfl

p

>

1<

: 9 fafltfl.

•

12 34 56 7 89 12 34 56 7 89 1 2 34 56 7 89

Figure 2-27:Quantification of CPD repair by photolyase

Quantification of CPD repair by photolyase of Fig. 2-25. Photoreactivation of clusters 1 to 9 was

quantified in DNA (DNA, grey bars), nucleosomes (empty bars) and in nucleosomes incubated with

yISW2 (dash bars) and ATP (black bars). Repair is shown as the fraction of CPD removed by 15, 30

and 60 minutes ofphotoreactivation.

60

2. Results

0 10 20 30 40 50 60 0 10 20 30 40 50 60

time [min] time [min]

Nucl. + ISW2 Nucl. + ISW2 + ATP

100 i 1 i

0 10 20 30 40 50 60 0 10 20 30 40 50 60

time [min] time [min]

Figure 2-28:Quantification of CPD repair by photolyase

Quantification ofCPD repair by photolyase of Fig. 2-25. Photoreactivation of clusters 1 to 9 was

quantified in naked DNA (DNA), nucleosomes (Nucl.) and in nucleosomes incubated with

yISW2 (Nucl.+ISW2) and ATP (Nucl.+ISW2+ATP). Repair of each cluster is plotted as function

of incubation time.

61

2. Results

DNA

Nucl.

ISW2

Nucl.

ISW2

ATP

•aa.

100

50

0

100

Nucl. & 50

0-10

100

.aCDa.CO

50

0

-10

100

50

15» 30' 60'

m_o_ nnn orüj

JEi

,

,

_ _;

Il : nH

kfl

.-JJJ liilL10 1214161811 1315 17 19

10 1214161811 1315 17 19

JjJltL10 121416 1811 1315 17 19

Figure 2-29:Quantification of CPD repair by photolyase

Quantification of CPD repair by photolyase of Fig. 2-26. Photoreactivation of clusters 10 to 19

was quantified in DNA (DNA, grey bars), nucleosomes (empty bars) and in nucleosomes

incubated with yISW2 (dash bars) and ATP (black bars). Repair is shown as the fraction of CPD

removed by 15, 30 and 60 minutes of photoreactivation.

62

2. Results

DNA Nucl.

+19^-18+17

-0-16

-e-15-gh14+ 13

-»«-12--11-•-10

0 10 20 30 40

time [min]

50 60 0 10 20 30 40

time [min]

50 60

Nucl. + ISW2 Nucl. + ISW2 + ATP

20 30 40

time [min]

0 10 20 30 40

time [min]

60

Figure 2-30:Quantification of CPD repair by photolyase

Quantification of CPD repair by photolyase of Fig. 2-26. Photoreactivation of clusters 10 to 19

was quantified in naked DNA (DNA), nucleosomes (Nucl.) and in nucleosomes incubated with

yISW2 (Nucl.+ISW2) and ATP (Nucl.+ISW2+ATP). Repair of each cluster is plotted as function

of incubation time.

63

2. Results

2.3.2. Effect of yISW2 on ATDED-long Nucleosomes

In contrast to the ySWI/SNF complex, very little is known about the effect of yISW2

remodeling on nucleosomes in vitro. Therefore, several experiments were performed in an

attempt to characterize the activity of ISW2 on ATDED-long nucleosomes.

2.3.2.1. DNasel Footprinting

To gain information about the positions of nucleosomes after incubation with yISW2 and

ATP, DNasel footprint experiments were performed. ATDED-long nucleosomes labeled on either

strand were incubated with yISW2 either in the absence or in the presence of ImM ATP. In these

experiments, the nucleosome concentration was 10 ng/pl and the molar ratio was 0.8 yISW2 per

nucleosome. Nucleoprotein gels showed that >80% of the labeled DNA was folded into

nucleosomes, which migrated predominantly as one band (Nl; Fig. 2-31A and Fig. 2-32A, lanes

2). The mobility of the nucleosomes was reduced after incubation with yISW2 in the presence of

ATP (N2; lanes 4), compared to the mobility of nucleosomes incubated in the absence of ATP

(lanes 3) and ofyISW2 (lanes 2) with either strand labeled, as previously observed (Fig. 2-24).

The DNasel cutting pattern observed in nucleosomes and naked DNA (Fig. 2-3 IB and

Fig. 2-32B, compare lanes 2 and 3 with lanes 8 and 9) was similar to the pattern observed

previously (Fig. 2-3 and Fig. 2-4). Addition of yISW2 to naked DNA did not alter the DNasel

cleavage pattern neither in the absence (Fig. 2-3 IB and Fig. 2-32B, lanes 10 and 11) nor in the

presence ofATP (lanes 12 and 13). In nucleosomes, addition of the yISW2 complex without ATP

did not change the DNasel cleavage pattern (lanes 4 and 5).

On the top strand of the nucleosomes, we observed changes of DNasel accessibility in

presence of yISW2 and ATP, (Fig. 2-3 IB, lanes 6 and 7). The yISW2 activity led to additional

cleavage at MU 1364 and to enhanced cleavage between MU 1478 and 1496. Additionally, a

cutting pattern containing both nucleosomal and naked DNA site was observed between MU

1488 and 1420. These results are indicative of changes in the accessibility of DNA to DNasel

upon incubation with yISW2.

On the bottom strand, no obvious changes were observed after incubation with yISW2 and

ATP (Fig. 2-32B, lanes 6 and 7). yISW2 was active, since the nucleosomes incubated with

yISW2 and ATP which were used as substrates for the DNasel footprint had a reduced mobility

in native nucleoprotein gel electrophoresis (Fig. 2-32A, lane 4).

64

2 Results

A.

<

o

- + + yISW2- - + ATP

DNA

+ + yISW2- + ATP

DNasel

m^ km s-N2-NI

-D

1436

14581

14661

. £#$. ^V #% #!*& ^fe*?^

- für ztz Sa ,„s»

ï -«ill*n

^ S SI S É i

* »i, . «« «* »S,

„.D SBj|||ig

1478 «i S-«1111° *ïïï5a^B a«»- * ^s* #&m a*»,

-"

1488*S

i

St. SSS1

1496§«S *•*

1 3 5 7 9 11 13

2 4 6 8 10 12

Figure 2-31:DNaseI footprint of nucleosomes remodeled by yISW2A. Nucleoprotein gel analysis. ATDED-long DNA was labeled on the top strand (lane 1)

reconstimted into nucleosomes and incubated alone (lane 2), with yISW2 (lane 3) and 1 mM

ATP (lane 4) for 30' at 30°C. The molecular ratio was of 0.8 yISW2 per nucleosome and the

nucleosome concentration was 10 ng/ul. Bands represent naked DNA (D), nucleosomes (Nl)and nucleosomes with reduced mobility (N2). B. DNasel footprinting of the remodeled

nucleosomes described in (A). Digestion of naked DNA (lanes 8 and 9), DNA incubated with

yISW2 (lanes 10 and 11) and 1 mM ATP (lanes 12 and 13), nucleosomes (lanes 2 and 3),nucleosomes incubated with yISW2 (lanes 4 and 5) and 1 mM ATP (lanes 6 and 7) with DNasel.

A/G (lane 1) is a sequencing marker. Black dots mark nucleosome-specific DNasel cutting sites;

numbers show map units. Empty dot mark additional, empty squares enhanced cleavage sites

after remodeling.

65

2 Results

A. B.

+ + yISW2- + ATP

*L

Nucl. DNA

+ + - + + yISW2+ - - + ATP

"r —äHB^i^Äi -UJM asei

pipit ^HK ^HHP Vît

É^Hft T»

1487

1463

1443

1422

12 3 4

13591* a*»« nfe

9 11 1310 12

Figure 2-32:DNaseI footprint of nucleosomes remodeled by yISW2A. Nucleoprotein gel analysis. ATDED-long DNA was labeled on the bottom strand (lane 1)reconstimted into nucleosomes and incubated alone (lane 2), with yISW2 (lane 3) and 1 mM

ATP (lane 4) for 30' at 30°C. The molecular ratio was of 0.8 yISW2 per nucleosome and the

nucleosome concentration was 10 ng/ul. Bands represent naked DNA (D), nucleosomes (Nl)and nucleosomes with reduced mobility (N2). B. DNasel footprinting of the remodeled

nucleosomes described in (A). Digestion of naked DNA (lanes 8 and 9), DNA incubated with

yISW2 (lanes 10 and 11) and 1 mM ATP (lanes 12 and 13), nucleosomes (lanes 2 and 3),

nucleosomes incubated with yISW2 (lanes 4 and 5) and 1 mM ATP (lanes 6 and 7) with

DNasel. A/G (lane 1) is a sequencing marker. Black dots mark nucleosome-specific DNasel

cutting sites; numbers show map units.

66

2 Results

Since the DNasel footprint results obtained with nucleosomes labeled on the bottom

strand differed from those obtained with nucleosomes labeled on the top strand, an additional

DNasel footprint experiment was performed. The yISW2 complex used in this experiment came

from a new batch. ATDED-long nucleosomes labeled on the bottom strand were incubated with

increasing amount of yISW2 resulting in molar ratios of 1.1, 1.5, 2.1, 2.9 and 4.4 yISW2 per

nucleosome. The nucleosome concentration was 10 ng/ul in all reactions. The incubation was

performed either in the presence or absence of 0.5 mM ATP for 30 minutes at 30°C prior to

DNasel digestion.

Native nucleoprotein gel electrophoresis showed that the reconstitution was efficient

(Fig. 2-33, lane 2) and that the nucleosomes were bound by yISW2 to form a higher molecular

weight complex which migrated prominently as a defined band (lanes 3 to 12). With the lowest

ISW2 to nucleosome ratio (lanes 3 and 4), the nucleosome band with reduced mobility described

previously (N2) was detected in the presence (lane 4) but not in the absence of ATP (lane 3),

suggesting that the ISW2 activity of the second purification might be comparable to the fist one.

.

0.45

^ "

11

Q fc - +

0.6 0.85

1.5 I 2.1

- +1 - +

1.2 1.8 ISW2[jig]2.9 4.4 ISW2:Nucl.

- + ATP

* |#w* * -W

My%^L ÉMÉ :^y ts#

w*-*- ^^w *--~" wßm*^**°

1 |t"<| Nucl-ISW2

äK^UJB

-N2

-NI

-D

1 2 3 5 6 7 8 9 10 11 12

Figure 2-33:Native nucleoprotein gel analysis of nucleosomes remodeled by yISW2

ATDED-long DNA was labeled on the bottom strand (lane 1) reconstituted into nucleosomes (lane

2), incubated with the indicated amount ofyISW2 (lanes 3, 5, 7, 9 and 11) and 1 mM ATP (lanes 4,

6, 8, 10 and 12) for 30' at 30°C. The nucleosome concentration was 10 ng/ul. Bands represent

naked DNA (D), nucleosomes (Nl), nucleosomes with reduced mobility (N2), nucleosome-yISW2

complex (Nucl.-ISW2) and the wells (W).

DNasel digestion was performed and the DNA analysed by denaturing gel electrophoresis

(Fig. 2-34 and Fig. 2-35). Addition of the yISW2 complex without ATP did not change the

DNasel cleavage pattern (lanes 4, 5, 8, 9, 12, 13, 16, 17, 20 and 21) of the nucleosomes

67

2. Results

independently of the amount of complex added, although one might have expected to see a

footprint of the yISW2 complex on the nucleosomes, given the define band observed by band

shift assay (Fig. 2-33).

%

Nucl.

0.45 0.6 0.85 1.2 1.8

1.1 1.5 2.1 2.9 4.4

+ - + - + - + - +

DNA

ISW2 [ug]ISW2:Nucl.

ATP

DNasel

1455-1443

1433-

1422-

1414- «•

1402- *• -

1391- •

1383-g»--

1372 - I» a*

1360-

#4-# — m

-1362

-1340

x*** ^

13 5

7 9 11 13 15 17 19 21 23 252

J4

36 8 10 12 14 16 18 20 22^24 26

Figure 2-34:DNaseI footprint of yISW2 remodeled nucleosomes

DNasel footprinting of the remodeled nucleosomes described in Fig. 2-33. Digestion of naked

DNA (lanes 24 to 26), nucleosomes (lanes 2 and 3), nucleosomes incubated with the indicated

amount ofyISW2 (lanes 4, 5, 8, 9, 12, 13, 16, 17,20 and 21) and 0.5 mM ATP (lanes 6, 7, 10, 11,

14, 15, 18, 19, 22 and 23) with DNasel. A/G (lane 1) is a sequencing marker. Black dots mark

nucleosome-specific DNasel cutting sites; empty dot mark additional cleavage sites; numbers

show map units.

In the presence of ylSW2 and ATP, the DNasel cutting pattern was significantly affected

(lanes 6, 7, 10, 11, 14, 15, 18, 19, 22 and 23). The 10 bps repeat pattern of ATDED-long

nucleosomes was replaced by a multitude ofcutting sites. The resulting cutting pattern was rather

68

2 Results

uniform over the sequence, compared to those of untreated nucleosomes and naked DNA. Close

inspection reveals that the resulting cutting pattern contains nucleosome and naked DNA specific

cutting sites as well as new cutting sites, for example at MU 1340.

Nucl. DNA

%

0.45 0.6 0.85 1.2 1.8

1.1 1.5 2.1 2.9 4.4

- + + + - + - +

ISW2 [jig]ISW2:Nucl.

ATP

DNasel

1497 — £/&* *M

I486

!2-S*-i Mi»*»-* "^m**

- üte Wtl I

LHk^ ate MK

— j<c as

r1455

1443 -3»

1433 ->1

1422-«•

1414-„•

1402-=*= =

J1391 - •

>23

«j* IS Si:MlaI aw^iIff*;

*. àÊâ 1

Ï@ï3i ,ss 5>*s ^*"

**w Ï8SÏ S8S ^^

»nm

4 6 1011121314151617181920212223242526

Figure 2-35:DNaseI footprint of yISW2 remodeled nucleosomes

DNasel footprinting of the remodeled nucleosomes described in (Fig. 2-33). The results of a long

run are shown. Description is as in Fig. 2-34.

2.3.2.2. Restriction Enzyme Analysis

The translational setting of nucleosomes remodeled by yISW2 was further investigated by

restriction enzyme analysis. Reconstituted nucleosomes labeled on the top strand were incubated

with yISW2 (first batch) at 30°C for 30 minutes and then subjected to digestion with the

restriction enzymes Pstl, Afllll, Hhal and Xhol, which cut 16, 31, 47/49 and 94 bps from the left

end of the DNA fragment, respectively (Fig. 2-1). The molar ratio was 0.8 yISW2 per

nucleosome and the nucleosome concentration was 20 ng/ul. On reconstituted fragments,

cleavage by Xhol was strongly inhibited (3% cut), in contrast to Pstl, Afllll and Hhal, which

restricted >80% of the DNA (Fig. 2-36), as observed previously (Fig. 2-5). No changes in the

cutting efficiencies were observed upon addition of yISW2 alone. In the presence of ATP,

cleavage by Pstl, Afllll and Xhol remained unchanged, whereas cleavage by Hhal dropped from

82% to 64%. It is not known whether Hhal, with a 4 bps recognition sequence, is inhibited on

69

2. Results

nucleosomes (Conconi et al., 1989). As a result of a movement of the histone octamer toward

more central positions on the ATDED-long DNA, the Hhal sites would be located at the left edge

of the newly positioned nucleosome. Thus, the drop of cutting efficiency observed in

nucleosomes after incubation with yISW2 and ATP might be consistent with a movement of the

histone octamer toward more central positions.

Pstl Afllll Hhal Xhol

+---+---+---+--- DNA- + + + -+ + + - +++- + + + Nucl.

-- + + -- + + --++--++ ISW2

---+---+---+---+ ATP

p^ NMm m/mrn,*"*** *öa£ä* ^^ W1P1ÜHP^HPf

MUM HEU 4tE& ^Ém**«****. -» màm., ^g^

f* ^ ^W ^^W WWW/w Hi

DPS- HPIf ^^^ ^ffl^

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16

%cut 98 86 85 86 J 99 88 88 88J100 82 82 64199 3 2 2

Figure 2-36:Restriction enzyme analysis of nucleosomes remodeled by yISW2

Digestion of naked DNA (lanes 1, 5, 9 and 13), reconstituted nucleosomes (lanes 2, 6, 10

and 14) incubated with yISW2 (lanes 3, 7, 11 and 15) and 0.5 mM ATP (lanes 4, 8, 12 and

16) with the indicated restriction enzymes. The cutting efficiencies are given in percent for

each lane (bottom).

2.3.2.3. UVPhotofootprinting

To further investigate the structure of nucleosomes after incubation with yISW2 and ATP,

a UV-footprint experiment was performed. Naked DNA was labeled on the top strand,

reconstituted in nucleosomes, incubated with yISW2 (first batch) either in the absence or

presence ofATP and irradiated with UV at a dose of 500 J/m2, yielding an average of 1 CPD per

fragment. Additionally, naked DNA was incubated with yISW2 either in the absence or presence

ofATP and irradiated with UV at a dose of 500 J/m2. In either case, a sample containing ATP but

without yISW2 was irradiated, since ATP absorbs UV light and therefore reduces the total

amount of CPDs for a given UV dose. The molar ratio was 0.7 yISW2 for both DNA and

nucleosomes.

Nucleoprotein gel analysis showed that nucleosome reconstitution was efficient and that

the nucleosomal products migrated predominantly as one band (Nl; Fig. 2-37, lane 11 and 12).

The nucleosomes were completely bound by yISW2 and appeared as a smear in the lane (lane

70

2. Results

15). After UV irradiation, most of the nucleosomes were relieved from yISW2 binding (lanes 18

and 19). In the presence ofATP, the nucleosomal band was retarded (N2; lane 19), as described

previously. In the naked DNA samples, a minor fraction of DNA was bound by yISW2 and

appeared as a smear (lane 4) which was reduced in the presence of ATP (lane 5) and after UV

irradiation (lanes 8 and 9).

DNA Nucl.

+ - - + + ISW2

+ - + - + ATP

- + + + + UV

-w

- - + + - - + + - -/+< - + - + - + - + < - +/-z Z s

Q - - - - + + + + Q fc - -

-N2-Nl

-D

12 3 4 5 6 7 8 9 11 10 12 13 14 15 16 17 18 19

Figure 2-37:Analysis of DNA and nucleosomes remodeled by yISW2 before and after

UV irradiation

Native nucleoprotein gel analysis. ATDED-long DNA was end-labeled on the top strand

(lanes 1 and 11), incubated alone (lane 2), with 0.5 mM ATP (lane 3), with yISW2 (lane 4)

and 0.5 mM ATP (lane 5). The end- labeled DNA was reconstimted in nucleosomes (lane 11),

which were incubated alone (lane 12), with 0.5 mM ATP (lane 13), with yISW2 (lane 13) and

0.5 mM ATP (lane 15). Unfortunately, the nucleosomes incubated with ATP and those

incubated with yISW2 were loaded in the same lane (lane 13). The samples described in lanes

3 to 5 and 12 to 15 were irradiated with UV at 500 J/m2 (lanes 6 to 9 and 16 to 19,

respectively). The molecular ratio of yISW2 to DNA and nucleosomes was of 0.7 for both

DNA and nucleosomes. Bands represent naked DNA (D), nucleosomes (Nl), nucleosomes

with reduced mobility (N2) and the wells (W).

The DNA was isolated and damage formation was analysed by digestion with T4-endoV

and gel electrophoresis (Fig. 2-38). In naked DNA, the CPD formation pattern did not change

upon addition ofATP (Fig. 2-38, lane 18) or yISW2 (lane 19) and ATP (lane 20). In nucleosomal

DNA, the CPD formation was enhanced in the T-tract of cluster 4 compared to the CPDs formed

in naked DNA (Fig. 2-38, lanes 17 and 21), indicating changes in the DNA structure upon

folding into nucleosomes, as observed previously (Fig. 2-6). No alteration of the CPD pattern

was observed in the T-tracts of clusters 2 and 3, consistent with the position of the nucleosomes.

Incubation with ATP (lane 22) or yISW2 (lane 23) did not change the CPD formation pattern of

nucleosomes.

71

2 Results

5'

DNA

- - + +

- + - +

+ + + +

Nucl.

- + +

+ - +

+ + + +

DNA

- - + +

- + - +

+ + + +

Nucl.

- - + +

- + - +

+ + + +

DNA

- - + +

- + - +

+ + + +

+ + + +

Nucl.

- - + + ISW2

- + - + ATP

+ + + + UV

+ + + + T4EndoV

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24

Figure 2-38:UV-footprint of DNA and nucleosomes remodeled by yISW2DNA of the samples described in Fig. 2-37 was isolated (lanes 9 to 16) and digested with T4-endoV

(lanes 1 to 8 and 17 to 24). Nucleosome and position of the damage clusters (to the left) and T-tracts

of clusters 2, 3 and 4 (to the right) are indicated.

After incubation with yISW2 and ATP, no obvious changes in the CPD formation were

observed (lane 24). Only a very faint increase in CPD formation were observed in the T-tracts of

clusters 3 and 2, compared with the damage formation in nucleosomal and naked DNA. Since we

do not know how the CPD formation pattern of those T-tracts would look like if they were folded

in nucleosomes, we are in the impossibility to drive conclusions from this observation. In the T-

tract of cluster 4, the CPD damage formation pattern remained similar to that of the nucleosomes,

indicating that this T-tract remained folded in nucleosomes after incubation with yISW2 and

ATP.

72

2. Results

2.4. The ATDED-short Nucleosome

Previous work showed that photoreactivation was inhibited on a nucleosome reconstituted

on a 134 bps long DNA fragment (HISAT nucleosome, (Schieferstein and Thoma, 1998)). Using

the ATDED-long nucleosome as substrate for photoreactivation, we observed that a nucleosome

positioned on a 226 bps long DNA fragment locally inhibits repair by photolyase. In contrast to

the repair pattern on the HISAT nucleosome, a modulation of repair was observed in the

ATDED-long nucleosome, the damages located toward the edges of the nucleosome being

repaired more efficiently than those located in the middle (Fig. 2-10 and Fig. 2-11). The cause for

the different repair pattern observed on the two nucleosomes might reside in the different DNA

sequences; in the reconstitution and photoreactivation protocols, which were modified; or in the

length of the DNA fragment. To investigate the influence of the fragment length on the

photoreactivation pattern, we constructed a shorter ATDED fragment and performed repair

experiments. The ATDED-short fragment matches the region where the ATDED-long

nucleosome has been mapped, covering the 136 bps from MU 1364 to 1500 of the sequence. It

contains polypyrimidine tracts on the bottom strand (Fig. 2-39) which allow CPD repair analysis.

(12) cTT5'

(13) ctcTTTTTTTT5'

5'_ __

(14) TTTTctct5'c^i —~>

12 13 14 15 16 1718 19

(15) cctccTT5'

(16) cccTTTTTcaatcS'

~r

MU 1400 MU 1500 <17)ctatccS

(18) TTc5'

(19) TTcTTgTTgTTcTTTTc5'

Figure 2-39:Schematic drawing of the ATDED-short nucleosome

Indicated are map units (MU), position of the radioactive end-label (asterisks) and

pyrimidine clusters on bottom strand (DNA sequences numbered from 12 to 19). The

ellipse denotes the location of the histone octamer on the DNA fragment. The position of

the nucleosome centre is indicated (black dot).

2.4.1. Characterization of the ATDED-short Nucleosome

The 136 bps long AfllllNael fragment was end-labeled with 32P on the bottom strand and

nucleosomes were reconstituted by histone octamer transfer from chicken erythrocyte core

particles. The reconstitution products were characterized by native nucleoprotein gel

73

2. Results

electrophoresis, DNasel and UV footprintings. Native nucleoprotein gel electrophoresis showed

that about 70% ofthe labeled DNA was folded into nucleosomes (Fig. 2-40). However, it should

be emphasized that, in the majority of the nucleoprotein gels, half or more of the reconstituted

ATDED-short DNA migrated like naked DNA (see Fig. 2-43A). Since DNasel and UV footprint

as well as analysis of photorepair (see below) support high reconstitution efficiencies, we assume

that the ATDED-short nucleosome was not stable during native gel electrophoresis. This would

also explain the smear observed between the naked and the nucleosomal bands in native gels

(Fig. 2-40 and Fig. 2-43 A).

Figure 2-40:Native nucleoprotein gel electrophoresis

(containing 10% glycerol)ATDED-short DNA (DNA) was reconstimted into

nucleosomes (Nucl.). Bands represent naked DNA (D) and

nucleosomes (N).

-N

-D

DNasel digests of the reconstituted ATDED-short nucleosomes produced the stereotypical

10 bps repeat pattern of rotationally positioned nucleosomes (Fig. 2-41, lanes 3 to 7), while

naked DNA digests exhibited no such pattern (lanes 8 to 11). Despite of the short length of the

fragment, the nucleosomal ladder did not extend over the entire sequence, since a cutting pattern

similar to naked DNA was observed at the very left end of the fragment. Although the

translational setting of the ATDED-short nucleosome was shifted in comparison to the position

of the ATDED-long nucleosome, both exhibit a similar rotational setting (compare the 10 bps

repeats in Fig. 2-41 and Fig. 2-4). Additionally, a strong nuclease hypersensitive site was

observed at MU 1424. The location of this hypersensitive site close to the 5'-end of the T-tracts

of cluster 16 is reminiscent of the HISAT nucleosome, which had a DNasel hypersensitive site in

the vicinity of the 5'-end of its T-tracts (Schieferstein and Thoma, 1996).

74

2 Results

Nucl. DNADNasel

-1381

f A •**•#*-***12 3456789 10 11

Figure 2-41:ATDED-short nucleosomes have a defined rotational settingDNasel footprint of ATDED-short Nucleosome labeled on the bottom strand.

Digestion of reconstimted nucleosomes (lanes 3 to 7) and naked DNA (lanes 8 to

11) with DNasel A/G (lane 1) and T (lane 2) are sequencing markers. Black dots

mark Nucleosome-specific DNasel cutting sites; numbers show map units. The

putative position of the histone octamer and the center of the nucleosome are drawn

schematically to the left of the gel.

CPD formation in sequences containing T-tracts is modulated by folding of DNA in

nucleosomes (Schieferstein and Thoma, 1996). Since ATDED-long reconstitution did modify the

CPD formation of the T-tracts of clusters 13 and 16 (see Fig. 2-6) and that the ATDED-short

sequence also contains these clusters (Fig. 2-39), we further investigated ATDED-short

nucleosomes by UV footprint analysis.

Either DNA or nucleosomes were irradiated with UV at a dose of 500 J/m, yielding an

average of one CPD per fragment. The DNA was isolated and damage formation analysed by

digestion with T4-endoV and gel electrophoresis (Fig. 2-42).

75

2 Results

In nucleosomal DNA, the CPD formation was altered in the T-tracts of clusters 13, 14 and

16 compared to the CPDs formed in naked DNA (lanes 2 and 4), indicating changes in the DNA

structure upon folding into nucleosomes, as observed in the ATDED-long nucleosome (Fig. 2-6).

Figure 2-42:UV footprint of the ATDED-

short nucleosome

ATDED-short DNA (lanes 1 and 2) or

nucleosomes (lanes 3 and 4) were irradiated

with UV-light at a dose of500 J/m2 and digestedwith T4-endoV (lanes 2 and 4). Nucleosome

and position of the damage clusters (to the left)

and T-tracts of clusters 13 and 16 (to the right)are indicated.

3'

Taken together, ATDED-short is a chromatin model substrate consisting of a nucleosome

reconstituted on a short DNA fragment. The polypyrimidine tracts are folded in the ATDED-

short nucleosome, which has a define rotational setting.

2.4.2. Photoreactivation Is Inhibited in ATDED-short Nucleosomes

In order to investigate the influence of the fragment length on CPD accessibility, repair

experiments were performed using irradiated ATDED-short nucleosomes. Irradiation of

nucleosomes with UV light at a dose of 500 J/m2 yielded an average of one CPD per fragment.

Irradiated nucleosomes were incubated with E. coli DNA photolyase at 30°C in the presence of

photoreactivating light for up to 120 minutes. Native nucleoprotein gel analysis showed that a

DNA

UV + +

T4EndoV - +

Nucl.

+ +

ZT5

T8

76

2 Results

large fraction (40-65%) of the DNA migrated as naked DNA (Fig. 2-43A, lanes 2 to 8). Similar

observations were made in several reconstitution experiments (Fig. 2-40 and not shown). Since

DNasel and UV footprintings (Fig. 2-41 and Fig. 2-42) as well as CPD repair analysis (see

below) support high reconstitution efficiencies, we assume that the nucleosome were

destabilized by nucleoprotein gel electrophoresis rather than during photoreactivation.

For repair analysis, UV-irradiated nucleosomes and DNA isolated from these irradiated

nucleosomes were treated with photolyase. The remaining CPDs were analysed (Fig. 2-43B) and

quantified in CPD clusters numbered from 12 to 19 (Fig. 2-39). The percentage of repair is

depicted either as function of the localisation of the clusters on the DNA sequence (Fig. 2-44 B)

or as function of the photoreactivation times (Fig. 2-44 C).

Comparison of the initial damage and the repair rates of the individual clusters shows no

correlation between the amount of damages and the repair efficiency (Fig. 2-44, A and B). CPD

repair in naked DNA was much faster than in nucleosomal DNA. In naked DNA, CPDs were

repaired to completion within 45 minutes (Fig. 2-43 B, lane 15). In addition, repair of all damage

clusters was rather homogeneous (Fig. 2-44, B and C).

In reconstituted DNA, repair was slow and inefficient. Repair of clusters 13 to 19, which

are folded into the nucleosome, remained poorly repaired (<50%) even after 60 minutes

photoreactivation (Fig. 2-43 B, lanes 6 to 11; Fig. 2-44, B). Only cluster 12 was repaired

efficiently (>90% after 30 minutes). Cluster 12 is at the very left end of the DNA fragment,

where the DNasel cutting pattern was identical to naked DNA (Fig. 2-41). Similarly, the repair

kinetics of cluster 12 was fast, while the clusters 13 to 19 had slow repair kinetics (Fig. 2-44 C).

The strong repair inhibition observed in nucleosomes supports efficient reconstitution and

stable nucleosomes, in contrast to the results obtained by nucleoprotein gel electrophoresis

(Fig. 2-43 A). We assume that the nucleosomes were not stable during nucleoprotein gel

electrophoresis (as discussed above), thus, we do not know how much naked DNA was present

during photoreactivation.

Our results indicate that the presence of a positioned nucleosome on the ATDED-short

DNA had an inhibitory effect on CPD repair by photolyase.

A modulation of repair was observed within the nucleosome. Clusters 13, 14, 15 and 19

were repaired more efficiently than clusters 16, 17 and 18 (Fig. 2-44 B and C). Although CPD

repair was modulated in both the ATDED-short and ATDED-long nucleosomes, repair of

individual clusters differed between the two nucleosomes. Notably, increased repair of clusters

13, 14 and 15 was observed in ATDED-short compared to ATDED-long. On the short fragment,

nucleosomes adopted a different position than on the long fragment, resulting in a shift of the

77

2 Results

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Figure 2-43:Photoreactivation ofATDED-short nucleosomes

A. Native nucleoprotein gel analysis (containing 10% glycerol). ATDED-short DNA (lane 1) was

reconstituted into nucleosomes, irradiated with 500 J/m2 (lane 2) and photoreactivated for 5, 15, 30,

45, 60 and 120 minutes (lanes 3 to 8, respectively). Bands represent naked DNA (D) and

nucleosomes (N). B. Photoreactivation of the samples described in (A). The results of a short run (to

the left) and a long run (to the right) are shown. UV-irradiated nucleosomes (Nucl.) and DNA

isolated from these nucleosomes (DNA) were treated with photolyase for the indicated times (min.).

Lane 4, initial CPD distribution; lanes 6 to 11, photoreactivation in nucleosomes; lanes 12 to 15,

photoreactivation in naked DNA. DNA isolated from non-irradiated nucleosomes (lanes 1 and 2),

undigested DNA (lane 3) and overdigested DNA (lane 5) are controls. The position of the histone

octamer and the center of the nucleosome are drawn schematically to the left ofthe gels. The damage

clusters that were quantified (see Fig. 2-44) are indicated (numbered from 12 to 19).

78

2. Results

Initial CPDs

0.2

Qa.

S 0.1

n nlilJ

B.

DNA

Nucl.

120

50

0

120

50

c.

a

a 60

12141618

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15* 30' 45760*

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dyade

DNA

121416 181315 17 19

4dyade

121416181315 17 19

dyade

Nucl.

20 30

time [min]

20 40 60 80 100 120

time [min]

Figure 2-44:Quantification of photoreactivation in ATDED-short nucleosomes

A. Initial damage of clusters 12 to 19. B. Photoreactivation of clusters 12 to 19 was

quantified in DNA (DNA, empty bars) and nucleosomes (Nucl., black bars). Repair is

shown as the fraction of CPD removed by 15, 30 and 45 minutes of photoreactivation for

naked DNA and by 15, 30 and 60 min. of photoreactivation for nucleosomes. The positionof the putative dyade is indicated. C. Photoreactivation of clusters 12 to 19 was quantifiedin DNA (DNA) and nucleosomes (Nucl.). Repair is plotted against photoreactivation time

for each cluster.

79

2 Results

damage cluster with respect to the octamer position and possibly explaining the differences in

repair of individual clusters. In addition, some naked DNA might be present in the ATDED-short

nucleosomes sample and contribute to the repair pattern.

80

3. Discussion

3. Discussion

DNA accessibility in eukaryotes is affected by intrinsic properties of nucleosomes, their

higher order organisation, and by chromatin modifying activities. Here, we studied whether

chromatin remodeling factors could overcome the inhibitory effects of chromatin on DNA repair.

Neither DNA-damage formation by UV-light nor interaction with photolyase disrupted a

preformed nucleosome in vitro. However, two different chromatin remodeling activities were

shown to remodel UV damaged nucleosomes and facilitate DNA-repair. Thus, it is possible that

remodeling activities might work in vivo to assist damage recognition and repair in chromatin

substrates.

3.1. CPD Formation and Repair in ATDED Nucleosomes

Fragments of the ATDED sequence (136 and 226 bps) were used to reconstitute

nucleosomes. While the short 136 bps sequence forces DNA winding on a histone octamer, the

long 226 bps sequence provides space for multiple positions and/or octamer movements. DNasel

footprinting, restriction enzyme accessibility assay and UV footprinting were used to assay the

rotational and translational settings of the reconstituted nucleosomes. Nucleosomes reconstituted

on the 226 bps long ATDED-long sequence are positioned toward the right end of the DNA

fragment, leaving the left end free. Thus, reconstitution ofATDED-long DNA provides a define

substrate containing both naked and nucleosomal DNA in the same complex. Upon nucleosome

reconstitution, some of the T-tracts (clusters 12 to 19 on the bottom strand and cluster 4 on the

top strand) are folded into nucleosomes while others are on the protruding free DNA (clusters 2

to 3 on the top strand), allowing the analysis of CPD formation and repair at various sites.

Our results indicate that the ATDED-long nucleosome is positioned on the DNA, however,

we cannot rule out that a fraction of the octamers might adopt alternative positions on the

fragment. The 136 bps ATDED-short nucleosome allows us to assess the influence of the

fragment length on nucleosome behaviour. DNasel footprinting showed that the ATDED-short

nucleosome had a define rotational setting. Despite of the shortness of the DNA fragment,

ATDED-short nucleosomes were not positioned centrally on the fragment, since about 10-15 bps

behaved as naked DNA at the very left end. Similar observations have been made previously

with nucleosomes reconstituted on fragments of the 5 S rDNA genes that contained a truncated

positioning sequence (Dong et al., 1990).

81

3. Discussion

The rotational setting of nucleosomes reconstituted on the short and on the long fragments

were very similar, consistent with the idea that DNA sequences have a preference to maintain

their rotational setting, while the translational setting is less enforced (Buttinelli et al., 1993;

Flaus et al., 1996; Pennings et al., 1991).

3.1.1. UV-Damage Formation and Nucleosome Stability

UV irradiation induces DNA lesions which distort the DNA-structure. Therefore, DNA

lesions may interfere with the stability of nucleosomes, promote their disruption or induce

sliding to alternate positions. The currently available data are somewhat controversial. Generally,

UV-damaged nucleosomes appear to be quite stable, since they can be purified from irradiated

cells (e.g. (Gale et al., 1987)). On the other hand, irradiation destabilized nucleosomes

reconstituted on 5S-rDNA (Mann et al., 1997) and plasmids (Matsumoto et al., 1994), while

nucleosomes reconstituted on HISAT DNA and 5S-rDNA were not affected (Liu et al., 2000;

Schieferstein and Thoma, 1996; Schieferstein and Thoma, 1998). In the examples shown in this

study, UV-lesions were not sufficient to disrupt the nucleosomes. Thus, structural distortions

introduced by UV-lesions were accommodated by both nucleosomes and nucleosome

remodeling complexes. This is consistent with the observation that crystallised nucleosomes can

accept the deficit of a base pair in one turn of the superhelix (Luger et al., 1997) and therefore

could as well accept a DNA-lesion.

3.1.2. Nucleosomes Inhibit Photoreactivation

Previous work reported a strong inhibition of CPD repair by photolyase and T4-

endonuclease V in nucleosomes in vitro as well as some site specific repair on the nucleosome

surface (Kosmoski et al., 2001; Schieferstein and Thoma, 1998). In reconstituted ATDED

nucleosomes, photoreactivation was severely inhibited in nucleosomal DNA, while repair in

naked DNA was fast. The inhibition of photoreactivation was restricted to the nucleosome, since

naked DNA protruding out of the nucleosome was repaired efficiently on the same substrate.

Thus, it appears that binding and/or processing of CPDs is strongly inhibited on the nucleosome

surface.

Photolyase is thought to specifically bind CPDs by flipping the dimer out of the DNA

duplex into the active site of the enzyme (Park et al., 1995; Vande Berg and Sancar, 1998). T4-

endonuclease V also uses a flip-out mechanism to cut DNA selectively at CPDs and has an

activity similar to photolyase on nucleosomes in vitro (Schieferstein and Thoma, 1998). Such a

flip-out mechanism implies that additional distortions of DNA in the vicinity of the lesion must

be induced by photolyase binding. Despite of the flexibility of nucleosomal DNA with respect to

82

3. Discussion

damage accommodation, nucleosomal DNA might not tolerate such distortions and therefore

exert a strong inhibition on photoreactivation. Consequently, efficient repair of nucleosomal

DNA requires disruption or displacement of nucleosomes with or without the help of remodelmg

activities.

More detailed analysis revealed that photoreactivation was modulated in the ATDED-long

nucleosome. Damages located at more distal regions of the nucleosome were repaired faster than

those located near the center. Enhanced DNA accessibility toward the edges compared with the

center of nucleosomes were observed with restriction enzymes and DNA binding factors (Polach

and Widom, 1995; Studitsky et al., 1997; Vettese-Dadey et al., 1994). In addition, instability of

the DNA termini of the nucleosome core was observed in melting and ionic strength-dependent

dissociation studies (McGhee and Felsenfeld, 1980; McGhee et al., 1980; Seligy and Poon,

1978). It has been proposed that the DNA might dissociate from the nucleosome at the entry and

exit points of the DNA and eventually proceed into the histone octamer and reassociate, thus

generating a dynamic equilibrium. According to this model, CPDs near the edges ofnucleosomes

should be more accessible than CPDs near the center. Alternatively, short-range movements of

the octamer on the DNA or the presence of alternate nucleosome positions could give rise to the

observed CPD repair pattern.

Photoreactivation of the ATDED-short nucleosome was also modulated, although to a

lesser extend than in the ATDED-long nucleosome. The differences in the repair pattern of both

substrates may be due to the distinct features of the nucleosomes, since ATDED-short and

ATDED-long nucleosomes appeared to adopt different positions on the DNA sequence.

Additionally, ATDED-short was less stable than ATDED-long nucleosome. Since ATDED-short

nucleosomes appeared to fall apart during native gel electrophoresis (see Section 2.4.1.), the

amount of free DNA present in the reaction and its contribution to the repair pattern cannot be

determined. The presence of naked DNA in the reaction would have a smothering effect on the

repair pattern, in particular for the damages which are not repaired in nucleosomes.

In the ATDED-long nucleosome, the lesions located on the 'linker side' of the nucleosome

were repaired more efficiently than those located at the other edge. Interestingly, enhanced

accessibility for DNasel was also observed around these sites. The presence of 'linker DNA' may

provide a structural flexibility in neighbouring nucleosomal DNA, resulting in enhanced

accessibility. Alternatively, ATDED-long nucleosomes might harbour a particular DNA structure

at these sites. For example, the numerous T-tracts of the ATDED sequence could contribute to

generate DNA structures which differ from canonical nucleosomal DNA. It is unlikely that a loss

83

3. Discussion

of histones was responsible for this observation since the mobility of the nucleosomes on native

nucleoprotein gels remained constant during photoreactivation. However, it remains to be

determined whether the nucleoprotein gels used are sensitive enough to detect the lost of a H2A/

H2B dimer.

In contrast to DNase I, which binds to the minor groove and generates single strand cuts,

photolyase bends DNA and flips out the pyrimidine-dimer into its active site (Park et al., 1995;

Vande Berg and Sancar, 1998). No obvious correlation of site specific photoreactivation with the

rotational setting as determined by DNase I was observed in ATDED nucleosomes. Moreover,

the 5'-end of damage cluster 16 of the ATDED-short nucleosome, which contains a nuclease

hypersensitive site (MU 1424) was poorly repaired, indicating that the requirements for DNasel

and photolyase accessibility are different.

3.1.3. Photolyase Does not Induce Octamer Movements on ATDED-

long Nucleosomes

Previous work showed that temperatures higher than 27°C induce short-range nucleosome

sliding on the 5 S rDNA sequence at low ionic strength (Meersseman et al., 1992). The multiple

positions adopted by nucleosomes reconstituted on 146 bps long fragments containing the 5 S

rDNA sequence are transformed to the central positions by heating to 37°C for 40 minutes at low

ionic strength. However, heating nucleosomes reconstituted on 180 bps long fragments

containing the 5 S rDNA sequence in similar conditions did not induce any nucleosome sliding

(Flaus et al., 1996). On the other hand, heat-induced sliding was observed in nucleosomes

reconstituted on DNA fragments of various length derived from the mouse mammary tumor 3'

long terminal repeat (MMTV 3'LTR) DNA (Flaus and Richmond, 1998). Thus, it appears that

the sequence on which nucleosomes are reconstituted has a significant influence on their stability

and on the ability of the octamer to slide along the sequence.

Despite of the length of the ATDED-long fragment, which would have allowed for

octamer movement along the DNA, both the repair pattern and DNasel footprint of reconstituted

nucleosomes showed that the histone octamer was not displaced during photoreactivation, which

was performed at 30°C for up to two hours. In addition, no changes in the translational setting of

ATDED-long nucleosomes were observed by analysis of photoreactivation samples in native

nucleoprotein gel lacking glycerol, which separate the different nucleosome positions on a DNA

fragment (Meersseman et al., 1992).

84

3. Discussion

DNA sequence is a major determinant of nucleosome positioning. It is energetically

favourable to accommodate a DNA sequence that is already curved around a histone octamer, as

DNA has to be bent around the histone core anyway (Drew and Travers, 1985a; Shrader and

Crothers, 1989). Naturally occurring nucleosome positioning sequences include the MMTV

promoter, a satellite DNA sequences and the 5 S ribosomal RNA gene (for reviews see

(Simpson, 1991; Thoma, 1992)). The commonly used 5S rDNA sequence positions nucleosomes

rotationally. However, nucleosomes reconstituted on the 5 S rDNA sequence adopt multiple

translational settings (Buttinelli et al., 1993; Dong et al., 1990; Pennings et al., 1991). Since

nucleosomes reconstituted on the ATDED sequence appear to adopt a preferential translational

position, ATDED represent an alternative nucleosomal substrate for in vitro approaches.

A particularity of the ATDED sequence is its numerous T-tracts of various length. The

presence of T-tracts might contribute to stabilize the octamer on the right half of the DNA

fragment. For example, nucleosome reconstitution on DNA containing a 20 nucleotide long T-

tract showed a strong preference of the T-tract for the edge of the nucleosomal particle (Prunell,

1982). Since the T-tracts of the ATDED sequence have been previously shown to form

nucleosomes in vitro (Losa et al., 1990), the ATDED-long sequence apparently contains a

nucleosome positioning sequence which prevents octamer movements on the fragment. Such a

positioning signal has been postulated before for the ATDED-long sequence (Losa et al., 1990).

This signal could be related to the position of the first nucleosome of the DED1 gene, since the

sequence from approximately MU 1434 towards the right end of the ATDED-long segment

corresponds to half of the sequence of the first nucleosome of the DED1 gene mapped in vivo.

3.1.4. Nucleosomal Inhibition of Photorepair Cannot Be Overcome

by Sequential Addition of Photolyase

Limiting factors in enzymatic reactions include the substrate to enzyme ratio and the

enzyme stability. In our experiments, the amount of photolyase used was sufficient for the

complete repair of naked DNA indicating that sufficient amount of repair enzyme were applied.

Since the repair of nucleosomal DNA is slow and inefficient, photolyase stability might be the

rate-limiting step in repair of nucleosomes. To test the stability of photolyase during our

photoreactivation experiments, fresh photolyase was added after 45 and 90 minutes incubation.

The general repair pattern of ATDED-long nucleosomes did not change. Since sequential

addition of fresh photolyase was not sufficient to achieve complete repair of nucleosomal CPDs,

the photolyase activity was apparently not the limiting factor in our experiments.

85

3 Discussion

Although the repair curves of some damage clusters (clusters 16, 17 and 18) were not

influenced by the addition of fresh photolyase, repair of some clusters (clusters 13, 14, 15, and

19) increased. The observation that the four most distal clusters were affected by additional

photolyase suggests that they might be transiently more accessible to photolyase. Several

possibilities exist which could explain a transient accessibility of these lesions. Nucleosome

wobbling within a few base pairs would transiently change the accessibility of nucleotides on the

nucleosome surface (Tanaka et al., 1996). However, the entire nucleosome would be affected and

not only clusters 13, 14, 15 and 19. Alternatively, sliding of the histone octamer on the DNA

fragment would presumably increase the accessibility of damages located at distal positions on

the nucleosome, while lesions located toward the center would remain unaffected. Two

observations argue against this explanation. The first one is that translational movements of the

histone octamer on the DNA fragment were not observed by native nucleoprotein gel

electrophoresis, the second that only about 20 bps of DNA are available on the right side of the

ATDED-long nucleosome, and that a shift of 20 bps would not last to free clusters 14 and 15

from being folded into nucleosomes. However, we cannot exclude that octamer movements

might occur transiently, thus remaining undetected by nucleoprotein gel analysis and that the

octamer might slide 'off the end' of the DNA fragment. Since clusters 13, 14, 15 and 19 are

located toward the edges of the ATDED-long nucleosome, another explanation may reside in the

differential accessibility of internal and terminal DNA in nucleosomes discussed above.

3.2. Remodeling by ySWI/SNF Alters the Structure of

Nucleosomal DNA and Facilitates CPD Accessibility

ySWI/SNF is the founding member of the swi2-family of ATP-dependent remodeling

complexes. It is 2 MDa in size and has been shown to increase the accessibility of nucleosomal

DNA to transcription factors and nucleases in vitro (Cote et al., 1994; Logie and Peterson, 1997;

Owen-Hughes et al., 1996; Utley et al., 1997) and to induce octamer sliding (Jaskelioff et al.,

2000; Whitehouse et al., 1999).

The DNasel footprint of ATDED-long nucleosomes remodeled by ySWI/SNF resembles

more the naked DNA pattern than the nucleosomal pattern, in agreement with previous DNasel

footprinting studies (Cote et al., 1998; Cote et al., 1994; Owen-Hughes et al., 1996). Binding of

the ySWI/SNF complex in the absence of ATP did not alter the 10 bps ladder of ATDED-long

nucleosomes, although the amount of ySWI/SNF used was sufficient to shift the nucleosomes

completely in a nucleoprotein gel. A weak protection was observed between MU 1374 to 1383

86

3 Discussion

compared to control nucleosomes, suggesting that ySWI/SNF binding might alter DNA

accessibility at these sites. However, additional footprinting and cross-linking studies would be

required to substantial this observation.

UV-photofootprinting provides information about the structure of DNA, since the

formation of UV lesions is dependent on local DNA structure (Becker and Wang, 1984; Becker

and Wang, 1989b; Lyamichev, 1991). In nucleosomes, the CPD formation pattern, in particular

in the T-tracts, was changed compared to naked DNA. Folding of DNA in nucleosomes altered

the T-tracts structure, indicating that nucleosomal constraints dominate over those of the T-tracts,

as observed previously (Schieferstein and Thoma, 1996). Interestingly, binding of ySWI/SNF

alone had no obvious effect on the CPD pattern, indicating that the nucleosome remained intact.

Addition of ySWI/SNF and ATP changed the CPD formation pattern, which appeared similar to

that ofnaked DNA, indicating that the structural constraints in remodeled nucleosomes no longer

dominated over those of the T-tracts. However, since nucleosomes were recovered after

competing off the ySWI/SNF complex, we concluded that the CPD pattern is a result of altered

histone-DNA interactions rather than of histone dissociation. Thus, DNA appears to be relaxed

or extended enough in the nucleosome-ySWI/SNF-ATP complex to allow formation of the T-

tract structures.

A recent crosslinking study showed that direct DNA contacts are made by ySWI/SNF

upon nucleosome binding, and that significant changes occur in those contacts with DNA after

remodeling (Sengupta et al., 2001). Here, we observed that binding by ySWI/SNF in the absence

ofATP inhibits repair of damages located toward either edge of the nucleosome (clusters 12 and

19) whereas repair of damages located elsewhere was not significantly altered. Interestingly,

cluster 12 (MU 1376 to 1378) coincide with the region where a weak protection from DNasel

digestion was observed upon ySWI/SNF binding. These results suggest that the complex might

bind to a define site on nucleosomes. Of the damages protected after ySWI/SNF addition, those

located on the 'linker side' of the nucleosome were repaired efficiently after remodeling in the

presence of ATP, while those located at the other edge of the nucleosome remained poorly

repaired. Since nucleoprotein gel analysis showed that ySWI/SNF remained bound to the

nucleosomes during photoreactivation, these results suggest that the interactions of the complex

with DNA change after remodeling.

87

3. Discussion

After incubation of our model substrate with ySWI/SNF and ATP, repair of nucleosomal

DNA was substantially enhanced, and repair of 'linker' DNA slightly reduced. The net result

was a more uniform repair along the DNA fragment. Therefore, ySWI/SNF action appears to

eliminate the modulated repair pattern observed between free and nucleosomal DNA. This

observation extends previous findings of increased accessibility of nucleosome templates to

DNasel, restriction enzymes and transcription factors after SWI/SNF remodeling (Peterson,

2000; Vignali et al., 2000). Competition experiments performed after photoreactivation showed

that only a minor fraction of the nucleosomes were disrupted by ySWI/SNF activity, therefore,

we assume that repair of remodeled nucleosomes reflects an altered structure (complexed with

ySWI/SNF) and are not due to a loss of histones.

ySWI/SNF can remodel UV damaged nucleosomes and appears to generally facilitate the

accessibility of photolyase to nucleosomal DNA. Extensive destabilization of the nucleosomal

structure or transient octamer mobilization, eventually to the very ends of the DNA fragments,

could account for the observed repair pattern. Thus, ySWI/SNF remodeling appears to modify

the nucleosomal structure and/or octamer position to such an extent that the activity of a base

flip-out enzyme can be accommodated, thereby providing a mean by which the cell could

overcome the inhibitory effect of nucleosomes on photoreactivation.

3.3. Nucleosome Mobilization by yISW2 Influences CPD

Repair

The currently characterized ATP-dependent chromatin-remodeling complexes have been

divided into three groups based on the identity of their ATPase subunit: the SWI/SNF, the ISWI

and the Mi-2 groups (Boyer et al., 2000a). It has been proposed that complexes belonging to the

SWI/SNF and the ISWI groups use distinct mechanisms to remodel chromatin structure

(reviewed in (Peterson, 2000; Vignali et al., 2000)). ISWI containing complexes have been

shown to induce octamer sliding (Eberharter et al., 2001; Hamiche et al., 1999; Langst et al.,

1999). yISW2 is a two-subunits complex belonging to the ISWI group of ATP-dependent

remodeling factors which has been shown to influence nucleosome spacing in vitro (Tsukiyama

et al., 1999) and nucleosome positioning in vivo (Kent et al., 2001) which appear to be distinct

from remodeling activities ofthe SWI/SNF complex.

Here, we provide two lines of evidence that His-tagged-yISW2 can act on UV-damaged

nucleosomes and move the nucleosome from the end to a more central position in an ATP-

dependent manner: First, native nucleoprotein gel analysis suggested that yISW2 mobilizes the

88

3 Discussion

end-positioned ATDED-long nucleosomes to more central positions on the DNA fragment.

Second, the photoreactivation revealed an altered repair pattern, in particular, enhanced repair at

sites towards the end of the fragment and inhibition in the centre. Notably, the majority of

nucleosomes were not stably bound by yISW2 under our conditions since the complex was not

recovered by nucleoprotein gels. Thus, yISW2 remodeling activity appears to be similar to those

of the ISWI containing complexes, which have all been shown to induce octamer sliding in vitro

(reviewed in (Langst and Becker, 2001b)).

ISW2 remodeling, by increasing the mobility of nucleosomes, leads to an increased

accessibility of some lesions to photolyase and provides a way to achieve repair of nucleosomal

CPDs. A mobilization of the histone octamer in either direction would results in a photorepair

pattern resembling to the pattern observed in positioned nucleosomes of the yeast URA3 gene in

vivo (Suter et al., 2002). In the URA3 gene, photorepair towards the center of nucleosomes was

slower than at the edges. This finding let us to suggest that ISW2-like remodelers could facilitate

repair of nucleosomal lesions in vivo.

Previously, DNasel footprint experiment have been performed on mononucleosomes

remodeled by the ISWI-containing complexes NURF and CHRAC. NURF was shown to protect

some sites from cleavage and to create new cleavage sites on nucleosomes reconstituted on a 161

bps long fragment containing sequences upstream of the hsp70 promoter (Tsukiyama and Wu,

1995). However, since reconstituted nucleosomes adopt multiple positions on this sequence, no

correlation could be made between the nucleosome sliding activity of NURF and the DNasel

cleavage pattern. No changes in the DNasel cleavage pattern of nucleosomes reconstituted on a

146 bps fragment was observed after remodeling by CHRAC (Langst et al., 1999). Since the

nucleosome mobilisation activity of CHRAC was shown on nucleosomes reconstituted on 248

bps long fragments, it is not known whether CHRAC does not function on short nucleosomes, or

whether its activity cannot be detected by DNasel footprinting.

Here, DNasel footprintings performed using 0.5 yISW2 molecules per nucleosomes

indicated that the rotational setting of nucleosomal DNA was not completely lost after

remodeling by yISW2. On the top strand, remodeling by ISW2 resulted in changes in DNasel

cleavage toward the right half of the nucleosome, while the left half was not altered. This might

be consistent with translational movements ofthe histone octamer along the DNA. However, the

appearance of mixed pattern complicated the interpretation of the results. No obvious changes

were observed on the bottom strand under similar conditions. When DNase I footprint was

performed at higher ylSW2 to nucleosome molar ratios using a new batch of ylSW2, significant

changes in the cleavage pattern were observed on the bottom strand upon remodeling. The

89

3 Discussion

resulting cleavage pattern contained both nucleosomal and naked DNA specific sites, as well as

new cleavage sites. Such a pattern could result from octamer movements on the DNA, disruption

of histone-DNA contacts, histone dissociation, or any combination of those possibilities.

In a recent study, enhanced DNasel cleavage was claimed at one edge of reconstituted

nucleosomes upon binding of the drosophila ISWI ATPase in the absence of ATP (Langst and

Becker, 2001a). In our case, binding of the yISW2 complex in the absence ofATP did not alter

the 10 bps digestion ladder of ATDED-long nucleosomes, although the amount of yISW2 used

was sufficient to bandshift the nucleosomes completely in a nucleoprotein gel. Apparently

yISW2 binding did neither result in a bulk distortion of the nucleosome nor contact specific

DNA sites with a high affinity.

3.4. Is Chromatin Remodeling Involved in DNA Repair in

vivo?

The packaging of DNA into nucleosomes severely restricts the accessibility of DNA and

impedes a wide range of nuclear processes including CPD repair. To defend the cells against

extensive mutagenesis of the genome, all DNA-lesions need to be repaired efficiently

(Hoeijmakers, 2001b). Although there is a pronounced modulation by nucleosomes and other

protein-DNA interactions, both nucleotide excision repair and photolyase almost completely

remove UV-induced DNA-lesions (Smerdon and Conconi, 1999; Thoma, 1999). Thus, the

inhibitory effect of nucleosomes has to be overcome to allow damage recognition and

processing. High resolution repair analyses on positioned nucleosomes of the URA3 gene in

yeast showed similar repair patterns for NER and photolyase: fast repair in the linker and a

decrease towards the centre of the nucleosomes (Suter et al., 2002; Tijsterman et al., 1999;

Wellinger and Thoma, 1997). In combination with multiple positions found by nuclease

footprinting, those studies suggest that intrinsic mobility of nucleosomes might place a lesion in

the linker DNA, thus providing a window of accessibility for damage recognition (Suter et al.,

2002).

Alternatively, there is an increasing number of repair related proteins with potential roles

in chromatin remodeling (Green and Almouzni, 2002). CSB and its yeast homologue Rad26 both

belong to the SNF2 family and are involved in transcription coupled repair rather than in repair

of non transcribed DNA (de Boer and Hoeijmakers, 2000; Eisen et al., 1995). In addition to their

roles in DNA repair, CSB and RAD26 are involved in transcription elongation (Jansen et al.,

2000; Lee et al., 2001; Selby and Sancar, 1997; Tantin et al., 1997; Woudstra et al., 2002).

90

3. Discussion

Recombinant CSB was shown to remodel undamaged nucleosomes and nucleosome arrays in

vitro thus being the first repair enzyme with remodeling activity (Citterio et al., 2000a). Rad7-

Radl6 is a complex of the NER pathway of yeast S. cerevisiae which is essential for repair of

non-transcribed chromatin (Bang et al., 1992; Verhage et al., 1994) and recognises UV-lesions in

an ATP-dependent way in vitro (Guzder et al., 1998). Radio has homology to SNF2 and might

play a role in nucleosome remodeling to generate space for the other NER proteins (Eisen et al.,

1995; Thoma, 1999).

The INO80 complex is a potential candidate for a chromatin remodeler involved in DNA

repair, since mutant yeast strains show a UV sensitivity phenotype (Shen et al., 2000). It is not

known to date whether this phenotype is due to defects in DNA repair itself, or if the complex

regulates the expression of genes involved in repair pathways. ACF, on the other hand, is a

chromatin assembly and remodeling factor containing ISWI and Acfl, which was shown to

facilitate NER of a specific lesion located in linker DNA, but not of a lesion in the nucleosome

(Uraetal.,2001).

The observation made here that two different remodeling activities, ySWI/SNF and

yISW2 can act on damaged nucleosomes and alter the accessibility for a base-flip out enzyme

suggests that similar activities can indeed play a role in vivo. Since many different complexes

have been identified in a wide range of species, it is tempting to postulate that at least some of

them might have redundant functions which have remained undiscovered until now. In addition,

some homologous ATPases have not been characterized yet. A clear challenge for the future is to

identify the factors which facilitate DNA repair in chromatin of living cells.

Nucleosome remodeling complexes are recruited to the promoter regions of specific genes

by transcriptional factors to regulate transcription (reviewed in (Peterson and Logie, 2000)). The

situation is different for DNA repair, since DNA-lesions are generated all over the genome and

need to be removed everywhere. This implies either that specialised complexes capable of both

damage recognition and nucleosome remodeling are involved, or that DNA-lesions need to be

recognised first, before nucleosome remodeling activities can be recruited. In the later case,

damage accessibility depends only on the structural properties of the region containing the DNA-

lesion, (e.g. nucleosome, linker, nucleosome free region) and on the genetic activity, e.g. whether

it is transcribed or replicated. We might speculate about a more general role of remodeling

activities in chromatin organization. Since all ATP-dependent remodeling complexes apparently

can change the positions of nucleosomes, it seems conceivable that these complexes might also

act randomly on the chromatin substrate in order to enhance the intrinsic dynamic properties of

nucleosomes and keep chromatin in a 'fluid' state. This would facilitate any DNA-sequence

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3. Discussion

recognition, in transcriptional regulation and repair, and, in addition, adjust packaging

constraints imposed by chromosome metabolism. Thus, remodeling complexes might perform a

rather general role in maintenance of chromosome structure.

With our model system we have tested a possible contribution of nucleosomes and

remodeling activities towards DNA-repair. It will be important to investigate whether similar

observations can be made with the eukaryotic enzymes and base excision repair, and whether a

contribution of remodeling activities can be detected as a requirement or help for repair

processes in living cells.

3.5. ySWI/SNF and yISW2 Remodeling: Different

Mechanisms?

It has been postulated that members of the SWI/SNF and the ISWI families remodel

chromatin in different manners (reviewed in (Narlikar et al., 2002)). For example, the ATPase

activity of members of the SWI/SNF family is stimulated by both naked DNA and nucleosomes,

while members of the ISWI family are stimulated only by nucleosomes. Differences were also

observed in DNA extrusion assay (Havas et al., 2000). Whereas BRG1 and ySWI/SNF could

extrude cruciform DNA from inverted repeats of DNA and chromatin templates, ISWI could

perform this function only on chromatin templates. The hypothesis of mechanistic differences

between families ofremodelers is further supported by studies which showed that the ySWI/SNF

and yISW2 complexes have antagonistic effects on the transcription of the INOI gene in vivo

(Goldmark et al., 2000; Peterson et al., 1991; Pollard and Peterson, 1997; Sugiyama and Nikawa,

2001).

Although this study mainly focused on the influence of nucleosome remodeling on

photorepair, our results suggest that ySWI/SNF and yISW2 might use different mechanisms to

remodel chromatin. Photolyase might be used in the future to gain more information about the

remodeling mechanisms of both complexes. Repair analysis on remodeled nucleosomes after the

inactivation of the complexes, for example through competition or ATP depletion, might give

useful informations about the fate of the remodeled nucleosomes. Another series of experiments

would consist in investigating remodeling of the ATDED-short nucleosome. SWI/SNF was

shown to remodel nucleosomes reconstituted on 154 bps long fragment by transcription factor

binding and restriction enzyme cleavage assays (Cote et al., 1994; Jaskelioff et al., 2000).

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3. Discussion

Remodeling by ISW2 might require a longer DNA fragment to mobilize nucleosomes.

Therefore, comparison of both remodelers on a short nucleosome might help to discriminate

between their remodeling mechanisms.

93

4. Materials and Methods

4. Materials and Methods

4.1. ATDED Subcloning

4.1.1. pl8ATDED and pl8ATDED-c

The 212 bp HindllllBamHI fragment of p8ATDED (Losa et al., 1990) was gel purified,

filled in with T4 DNA polymerase (NEB), ethanol precipitated and ligated in either orientation

into the blunted Sad site of pUC18 to generate pl8ATDED and pl8ATDED-c. The ligation

products were transformed into E. coli XL-Blue competent cells. Positive clones were checked

by restriction analysis.

4.1.2. pGEM-ATDED-short

For subcloning of a short ATDED fragment, a fragment containing the ATDED-short

sequence and suitable restriction sites for subsequent fragment isolation and labeling was

generated using the ligation mediated PCR technique (Pfeifer et al., 1999; Pfeifer and Tornaletti,

1997).

The template for the PCR reaction was prepared as follows:

For the preparation of the linker, the primers Ulinker-ATsh and Llinker-ATsh (10 nmol

each) were mixed and the buffer adjusted to 250 mM TRIS pH 7.7 in a final volume of 500 pi.

The sample was incubated for 3 minutes at 95°C and the tube placed in a beaker containing 1.4 L

of 70°C warm water. The beaker was placed at 4°C for 3 hours. The annealed linker was stored at

-20°C and always thawed on ice.

p8ATDED (Losa et al., 1990) was digested with Xhol and BanI, generating several bands

including a 229 bp fragment containing the ATDED-short sequence. The digestion products were

purified by StrataClean (STRATAGENE) extraction and ethanol precipitation, and used as

template for one primer extension reaction with primer1-ATsh. 100 ng digested p8ATDED and

0.6 pmol primer 1-ATsh were mixed and the buffer adjusted to 40 mM TRIS pH 7.7 and 50 mM

NaCl in a final volume of 15 pi. The sample was incubated for 3 minutes at 95°C to denature the

DNA, followed by a 30 minute incubation at 45°C to allow primer annealing. The sample was

kept on ice and 7.5 pi of a freshly prepared mix containing 20 mM MgCl2,20 mM DTT and 0.25

mM of each dNTP was added. After the addition of 5 Units Sequenase 2.0 (USB), the reaction

94

4. Materials and Methods

mix was incubated for 10 minutes at 48°C and put on ice. 6 pi of 300 mM TRIS pH 7.7 was

added, the sample incubated at 67°C for 15 minutes to inactivate the Sequenase enzyme and kept

on ice.

100 pmol of linker was added to the sample, the buffer adjusted to 7.5 mM MgCl2, 17 mM

DTT, 0.9 mM ATP and 47 pg/pl BSA in a final volume of 80 pi. 3 Units T4 DNA ligase (USB)

were added and the reaction mix incubated at 18°C over night. The ligase was inactivated by

incubating the sample at 70°C for 10 minutes. 10 pg tRNA was added as carrier, the sample

ethanol precipitated and the pellet solved in 50 pi dH20.

For the PCR reaction, primer2-ATsh and primer3-ATsh (10 pmol each) were added to the

template and the buffer adjusted to 0.2 mM of each dNTP and lx PCR Buffer (SIGMA) in a final

volume of 100 pi. 3 Units Taq polymerase (SIGMA) were added and the PCR reaction

performed in a PERKIN ELMER 9600 thermocycler (25 cycles consisting of 1 minute at 95°C, 2

minutes at 55°C and 1.5 minute at 74°C). Since the tRNA used as carrier for the ethanol

precipitation migrated at the same height than the expected 162 bp amplification product, 1 pi of

the PCR products was used to perform a second PCR reaction using the same protocol. DNA was

purified over a Qiaquick PCR columns (QIAGEN), the 162 bp band gel purified and ligated into

the pGEM vector (Promega) to generate pGEM-ATDED-short. The ligation product were

transformed into E. coli DH5oc competent cells. Positive clones were checked by restriction

analysis and sequenced.

Table 4-1: Oligos for ligation mediated PCR

Name Sequence (5'-3')

Ulinker-ATsh TGCTTGGATCCTTAAGGCCTCTTGAAC

Llinker-ATsh GTTCAAGAGGC

primerl-ATsh CGATTACGAATTCGCCGGCAGTTCAGCCATAATATGA

primer2-ATsh TGCTTGGATCCTTAAGGCCTC

primer3-ATsh CGATTACGAATTCGCCGGCAG

4.2. Preparation of the DNA Fragments

4.2.1. Purification of the ATDED-long DNA Fragments

40 pg of plasmid DNA (pl8ATDED or pl8ATDED-c) were digested with 50 U Smal

(Roche) in 100 pi bufferA for 3h at 25°C. 50 U EcoRI (Roche) were then added and the samples

were incubated for 3h at 37°C. The DNAwas ethanol precipitated, solved in 64 pi TE pH 7.5 and

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4. Materials and Methods

loaded on a 0.8% low melting agarose gel. The gel was run without EtBr in 0.5 x TBE at 70 V

and 4°C for 2h30. The bands corresponding to the SmallEcoRI fragments were cut out of the gel

(without EtBr and without UV) and the DNA was isolated by AgarACE agarase (Promega)

digestion followed by ethanol precipitation. The DNA pellet was solved in TE pH 7.5 and its

concentration guessed by comparison with a known amount of marker DNA on agarose gels.

4.2.2. Purification of the ATDED-short DNA Fragment

40 pg of plasmid DNA (pGEM-ATDED-short) were digested with 50 U Aflll (NEB) and

Nael (Roche) in 100 pi buffer A supplemented with BSA for 3h at 37°C. The DNA was ethanol

precipitated, solved in 40 pi TE pH 7.5 and loaded on a 1% low melting agarose gel. The gel was

run without EtBr in 0.5 x TBE at 70 V at 4°C for 2h30. The 136 bp AfllllNael band was cut out

of the gel (without EtBr and without UV) and the DNA was isolated by AgarACE agarase

(Promega) digestion followed by ethanol precipitation. The DNA pellet was solved in TE pH 7.5

and its concentration guessed by comparison with a known amount of marker DNA on agarose

gels.

Table 4-2: Sequence of the ATDED fragments

Fragment name length Sequence (5'to 3')1'

ATDED 226 bp GGGTACCGAGCTTGGCTGCAGGTCATACGTGTCATTCTGAACGAG

GCGCGCTTTCCTTTTTTCTTTTTGCTTTTTCTTTTTTTTTCTCTTGAA

CTCGAGAAAAAAAATATAAAAGAGATGGAGGAACGGGAAAAAGT

TAGTTGTGGTGATAGGTGGCAAGTGGTATTCCGTAAGAACAACAA

GAAAAGCATTTCATATTATGGCTGAACTGAGGACGGATCCGAATT

ATDED-c 226 bp GGGTACCGGATCCGTCCTCAGTTCAGCCATAATATGAAATGCTTTT

CTTGTTGTTCTTACGGAATACCACTTGCCACCTATCACCACAACTA

ACTTTTTCCCGTTCCTCCATCTCTTTTATATTTTTTTTCTCGAGTTCA

AGAGAAAAAAAAAGAAAAAGCAAAAAGAAAAAAGGAAAGCGC

GCCTCGTTCAGAATGACACGTATGACCTGCAGCCAAGCTCGAATT

ATDED-short 136 bp GGCAGTTCAGCCATAATATGAAATGCTTTTCTTGTTGTTCTTACGG

AATACCACTTGCCACCTATCACCACAACTAACTTTTTCCCGTTCCT

CCATCTCTTTTATATTTTTTTTCTCGAGTTCAAGAGGCCTTAA

' the nucleotides added during labeling are in bold

4.2.3. End-labeling of the DNA Fragments

About 250 ng of purified fragments (Table 4-2) were incubated with 3 pi oc-[32P]-dATP

and 1.75 U Kleenow exo" (USB) at 30°C for 30 minutes in a final volume of 35 pi (0.5 mM

dTTP, lx Kleenow exo" buffer). To ensure that the fill-in reaction was complete, 10.5 pi dH20, 5

pi dNTPs (5 mM each), 2 pi 1 Ox Kleenow exo" buffer and 1.25 U Kleenow exo" were added to

96

4. Materials and Methods

the samples which were then further incubated for 15 minutes at 30°C. The reaction was stopped

by the addition of 15 pi 20 mM EDTA pH 7.5 and 1 pi StrataClean Resin (STRATAGENE). The

samples were vortexed and centrifuged in a microfuge at 13000 rpm for 3 minutes. The

supernatant was purified over a pre-equilibrated (TE pH 7.5) G-50 column (Roche) and

transferred to a new tube. The counts incorporated in 1 pi were determined using a scintillation

counter (model 1500 Tri-Carb, Camberra Packard) and the specific activity of the labeling was

extrapolated. Typically, we obtained a specific activity of 3-5 x 107 cpm/pg DNA.

4.3. Preparation of Nucleosome Core Particles

Nucleosome core particles depleted of histone HI and H5 were prepared from chicken

erythrocyte nuclei by U. Schieferstein (Schieferstein, 1997; Schieferstein and Thoma, 1998) and

stored at -80°C (2.1 mg/ml in 10 mM TRIS pH 7.5, 1 mM EDTA pH 7.5, 10 mM NaCl). To test

the integrity of the histones, 2.1 pg of cores were analysed by 18% SDS-PAGE electrophoresis

and Coomassie blue staining (Fig. 4-1).

cores

(2.1 ug)

Figure 4-1: SDS-PAGE analysis of histone proteins2.1 fig of nucleosome core particles were analysed on a

18% SDS-PAGE and stained with Coomassie blue. Bands

represent histones H3, H2B, H2A and H4.

4.4. Nucleosome Reconstitution

Reconstitution was done by histone octamer transfer from chicken erythrocytes core

particles as described (Schieferstein and Thoma, 1998) with modifications. 200 ng endlabeled

ATDED fragments were mixed with 8 pg core particles and 32 pi 5 M NaCl in a final volume of

200 pi (final concentration: 40 ng/pl cores, 1 ng/pl labeled DNA, 0.8 M NaCl, 8 mM TRIS pH

7.5 and 0.8 mM EDTA pH 7.5). The samples were incubated at 37°C for 30 minutes, at 4°C for

another 30 minutes and then dialysed against 1 liter of dialysis buffer 1 (0.6 M NaCl, 10 mM

97

4. Materials and Methods

TRIS pH 7.5, 1 mM EDTA pH 7.5 and 50 pM PMSF) in a PIERCE microdialyser (MWCO

10000) at 4°C over night. The buffer was changed for 1 liter dialysis buffer 2 (50 mM NaCl, 10

mM TRIS pH 7.5, 1 mM EDTA pH 7.5 and 50 pM PMSF) and the samples dialysed further at

4°C for 4-5 hours. The buffer was exchanged for 1 liter fresh dialysis buffer 2 and the samples

dialysed further for 2-3 hours at 4°C. The samples were recovered and the reconstitution

products analysed on nucleoprotein gels.

4.5. Remodeling with ySWI/SNF

ySWI/SNF was a kind gift of C. Peterson. The first batch contained 80 nM ySWI/SNF, 20

mM HEPES pH 8.0, 350 mM NaCl, 0.1% Tween 20, 1 mM DTT and 10% glycerol. The second

batch contained 130 nM ySWI/SNF, 20 mM HEPES pH 7.4, 350 mM NaCl, 0.1% Tween, 10%

glycerol and 200 pg/ml insulin.

For the DNasel footprint experiment, 10 pi ySWI/SNF (batch 1, 80 nM ySWI/SNF) was

added to 160 ng nucleosomes and the buffer was adjusted to 8 mM TRIS pH 7.5, 100 mM NaCl,

1 mM DTT, 5 mM MgCl2, 1 mM ATP, 3% glycerol in a final volume of 30 pi. The molar ratio of

ySWI/SNF to nucleosomes was 1 to 1 (10 ng complex/ng nucleosome). The samples were

incubated 30 minutes at 30°C. 3 pi were used for nucleoprotein gel analysis. The remaining 27 pi

were used for DNasel footprinting experiment.

For the UV photofootprint experiment, 5 pi ySWI/SNF (batch 1, 80 nM ySWI/SNF) was

added to 100 ng nucleosomes and the buffer was adjusted to 8 mM TRIS pH 7.5, 100 mM NaCl,

1 mM DTT, 5 mM MgCl2, 1 mM ATP, 3% glycerol in a final volume of 20 pi. The molar ratio of

ySWI/SNF to nucleosomes was 0.8 to 1 (8 ng complex/ng nucleosome). The samples were

incubated 30 minutes at 30°C. 1 pi was used for nucleoprotein gel analysis. Another 1 pi was

used for the competition experiment. The remaining 18 pi were used for UV footprinting

experiment.

For the photoreactivation experiment, 18.5 pi ySWI/SNF (batch 2, 130 nM ySWI/SNF)

was added to 800 ng nucleosomes and the buffer was adjusted to 8 mM TRIS pH 7.5, 100 mM

NaCl, 1 mM DTT, 5 mM MgCl2, 0.5 mM ATP, 3% glycerol, 46 pg/ml insulin in a final volume

of 80 pi. The molar ratio of ySWI/SNF to nucleosomes was 0.6 to 1 (6 ng complex/ng

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4. Materials and Methods

nucleosome). The samples were incubated 30 minutes at 30°C. 1.5 pi were used for

nucleoprotein gel analysis. Another 1.5 pi were used for the competition experiment. The

remaining 77 pi were used for photoreactivation experiment.

4.5.1. Competition of ySWI/SNF

1 pg linear plasmid DNA (pBSFT99 digested with Bgll in 10 mM TRIS pH 7.5, 100 mM

NaCl, 3% glycerol) was added to 1-1.5 pi of nucleosome-ySWI/SNF mixture, resulting in a final

volume of 5 pi. The reactions were incubated for 45 minutes at room temperature and analyzed

on nucleoprotein gels.

4.6. Remodeling with yISW2

His-tagged-yISW2 was a kind gift of D. Fitzgerald and T. Richmond. The first batch

contained 75 ng/pl yISW2 (270 nM), 15 mM TRIS pH 7.0,220 mM KCl, 0.03% NP-40, 0.3 mM

MgCl2, 30% glycerol and 100 pg/ml BSA. The second batch contained 500 ng/pl yISW2, 15

mM TRIS pH 7.0, 220 mM KCl, 0.03% NP-40, 0.3 mM MgCl2 and 30% glycerol.

For the photoreactivation experiments, 2.2 pg yISW2 (batch 1, 75 ng/pl yISW2) were

added to 2 pg nucleosomes or naked DNA and the buffer was adjusted to 10 mM TRIS pH 7.5,

90 mM NaCl, 5 mM MgCl2, 1 mM DTT, 0.5 mM ATP, 8% glycerol, 30 pg/ml BSA in a final

volume of 110 pi. The molar ratio of yISW2 to nucleosomes was 0.7 to 1 (1.1 ng complex/ng

nucleosome). The samples were incubated 30 minutes at 30°C. 3 pi were used for nucleoprotein

gel analysis. The remaining 107 pi were used for photoreactivation experiment.

For the DNasel footprint experiment, 330 ng yISW2 (batch 1, 75 ng/pl yISW2) were

added to 320 ng nucleosomes or naked DNA and the buffer was adjusted to 10 mM TRIS pH 7.5,

50 mM NaCl, 5 mM MgCl2, 1 mM DTT, 0.5 mM ATP, 4% glycerol, 13 pg/ml BSA in a final

volume of 35 pi. The molar ratio of yISW2 to nucleosomes was 0.7 to 1 (1 ng complex/ng

nucleosome). The samples were incubated 30 minutes at 30°C. 3 pi were used for nucleoprotein

gel analysis. The remaining 32 pi were used for DNasel footprinting experiment.

Alternatively, 0.45-1.8 pg yISW2 (batch 2, 500 ng/pl yISW2) were added to 300 ng

nucleosomes and the buffer was adjusted to 10 mM TRIS pH 7.5, 50 mM NaCl, 5 mM MgCl2, 1

mM DTT, 0.5 mM ATP, 4% glycerol in a final volume of 30 pi. The molar ratio of yISW2 to

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4. Materials and Methods

nucleosomes was 1.1-4.4 to 1. The samples were incubated 30 minutes at 30°C. 2 pi were used

for nucleoprotein gel analysis. The remaining 28 pi were used for DNasel footprinting

experiment.

For the restriction enzyme experiment, 1.5 pg yISW2 (batch 1, 75 ng/pl yISW2) were

added to 1.5 pg nucleosomes or naked DNA and the buffer was adjusted to 10 mM TRIS pH 7.5,

90 mM NaCl, 5 mM MgCl2, 1 mM DTT, 0.5 mM ATP, 9% glycerol, 30 pg/ml BSA in a final

volume of 75 pi. The molar ratio of yISW2 to nucleosomes was 0.7 to 1 (1 ng complex/ng

nucleosome). The samples were incubated 30 minutes at 30°C. 1.5 pi were used for

nucleoprotein gel analysis. The remaining 73.5 pi were used for restriction enzyme digestion

experiment.

For the UV photofootprint experiment, 1.5 pg yISW2 (batch 1, 75 ng/pl yISW2) were

added to 1.6 pg nucleosomes or naked DNA and the buffer was adjusted to 10 mM TRIS pH 7.5,

60 mM NaCl, 5 mM MgCl2, 1 mM DTT, 0.5 mM ATP, 6% glycerol, 20 pg/ml BSA in a final

volume of 100 pi. The molar ratio of yISW2 to nucleosomes was 0.7 to 1 (0.9 ng complex/ng

nucleosome). The samples were incubated 30 minutes at 30°C. 2 pi were used for nucleoprotein

gel analysis. The remaining 98 pi were used for UV footprinting experiment.

Alternatively, 0.8 or 2.5 pg yISW2 (batch 2, 500 ng/pl yISW2) were added to 1.3 pg

nucleosomes and the buffer was adjusted to 10 mM TRIS pH 7.5, 50 mM NaCl, 5 mM MgCl2, 1

mM DTT, 0.5 mM ATP, 2% glycerol in a final volume of 80 pi. The molar ratio of yISW2 to

nucleosomes was 0.5 or 1.4 to 1. The samples were incubated 30 minutes at 30°C. 2 pi were used

for nucleoprotein gel analysis. The remaining 78 pi were used for UV footprinting experiment.

4.7. DNasel Footprint

150-330 ng nucleosomes were incubated with 0.4 U DNasel (Roche; 0.4 U/pl in 10 mM

TRIS pH 8, stored at -20°C) in the presence of 5 mM MgCl2 at room temperature in final

volumes of 25-35 pi. Naked DNA samples were treated the same way, except that 0.04 U

DNasel were used. Aliquots were taken at various time points (45"-6'), the reaction stopped by

the addition of 10 pi 20 mM EDTA, vortexed and kept on ice until DNA purification. The

samples were digested with Proteinase K (Roche) at 50°C for 3 hours (final concentration: 0.5%

SDS, 0.5 mg/ml Proteinase K). To remove the digested proteins, 1.5 pi StrataClean Resin

(STRATAGENE) was added to the samples. The samples were vortexed and the volume adjusted

to 100 pi with H20. After centrifugation in a table top centrifuge at 13 000 rpm for 2 minutes, the

100

4. Materials and Methods

supernatant was recovered and transferred to a tube containing 10 pi NaAc pH 5.2. The DNA

was precipitated by adding 400 pi ethanol, vortexing, incubating the samples at -20°C for 1 hour

and centrifuging at 13 000 rpm and 4°C for another hour. The pellets were washed once with 100

pi cold 70% ethanol and dried in the speed-vac for 10 minutes. Scintillation counter (model 1500

Tri-Carb, Camberra Packard) was used to determine the counts contained in each pellet, which

were then solved in loading buffer (80% formamide, 10 mM NaOH, 1 mM EDTA, 0.1% Xylene

Cyanole, 0.1% Bromophenol blue) to have the same counts/pl in every samples of one

experiment (typically 2-7 x 103 cpm/pl). The samples were analysed on 8% denaturing

acrylamide gels.

4.7.1. Maxam-Gilbert Sequencing

End-labelled DNA was mixed with 6 pg salmon sperm DNA, ethanol precipitated, solved

in 14 pi dH20 and sequenced as previously described (Maxam and Gilbert, 1980; Sambrook et

al., 1989) with minor modifications. For the A/G reaction, 3 pi 8.8% formic acid were added to

the 14 pi DNA, incubated 7 minutes at 37°C and chilled to 0°C. For the T reaction, 9 pi DNA

was incubated for 2 minutes at 90°C and chilled to 0°C. 20 pi KMn04 (20 pg/ml) were added

and the samples incubated for 4 minutes at 20°C. 10 pi allyl alcohol were added and the samples

were vortexed for 30 seconds and lyophilized for 1 hour at 50°C in a speed vac.

150 pi fresh IM piperidine solution was added to the A/G and T reactions and the samples

incubated for 30 minutes at 90°C. 1.2 ml butanol were added, the samples vortexed and

centrifuged for 2 minutes at 13 000 rpm and 4°C. The supernatant was removed completely. 150

pi 1% SDS and 1.5 ml butanol were added to the pellet. After vortexing for 30 seconds, the

samples were centrifuged for 2 minutes at 13 000 rpm. The supernatant was removed completely,

the pellet washed with cold 70% ethanol and lyophilized for 20 minutes in a speed vac.

Scintillation counter (model 1500 Tri-Carb, Camberra Packard) was used to determine the counts

contained in each pellet, which were then solved in loading buffer (80% formamide, 10 mM

NaOH, 1 mM EDTA, 0.1% Xylene Cyanole, 0.1% Bromophenol blue) to have the same counts/

pi in every samples of one experiment (typically 2-7 x 103 cpm/pl). The samples were analysed

on 8% denaturing acrylamide gels.

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4 Materials and Methods

4.8. Restriction Enzyme Digestions

1.8 pg nucleosomes or naked DNA were incubated with 50 U of the restriction enzyme

Afllll, Hhal (NEB), Pstl or Xhol (Roche) at 30°C for 3 hours in a buffer adjusted to 100 mM

NaCl, 10 mM MgCl2, 1 mM DTT, 50 mM TRIS pH 7.5, and 0.1 mg/ml BSA for the restriction

enzymes Afllll and Hhal, in a final volume of 50 pi.

Alternatively, 250 ng nucleosomes remodeled by yISW2 were incubated with 15 U of the

restriction enzymes Afllll (NEB), Hhal, Xhol (Fermentas) or Pstl (Roche) at 30°C for 3 hours in

a buffer adjusted to 100 mM NaCl, 15 mM MgCl2, 1 mM DTT, 50 mM TRIS pH 7.5, and 0.1

mg/ml BSA for the restriction enzymes Afllll and Hhal, in a final volume of 25 pi.

The samples were digested with Proteinase K (Roche) (f.c. 0.5% SDS, 0.5-1.4 mg/ml

Proteinase K) for 3 hours at 50°C. To remove the digested proteins, 1-5 pi StrataClean Resin

(STRATAGENE) was added to the samples. The samples were vortexed and the volume adjusted

to 100 pi with H20. After centrifugation in a table top centrifuge at 13 000 rpm for 2 minutes, the

supernatant was recovered and transferred to a tube containing 10 pi NaAc pH 5.2. The DNA

was precipitated by adding 400 pi ethanol, vortexing, incubating the samples at -20°C for 1 hour

and centrifuging at 13 000 rpm and 4°C for another hour. The pellets were washed once with 100

pi cold 70% ethanol and dried in the speed-vac for 10 minutes. Scintillation counter (model 1500

Tri-Carb, Camberra Packard) was used to determine the counts of each pellet, which were then

solved in loading buffer (80% formamide, 10 mM NaOH, 1 mM EDTA, 0.1% Xylene Cyanole,

0.1% Bromophenol blue) to have the same counts/pl in every samples of one experiment. The

samples were analysed on 8% denaturing acrylamide gels.

4.9. UV Irradiation

Nucleosomes or naked DNA were irradiated in TE (pH 7.5), 50 mM NaCl or in

remodeling buffer at DNA concentration of 40 ng/pl or 5 ng/pl for the ySWI/SNF remodeled

samples. A petri dish was filled with ice and covered with parafilm. The samples were pipetted in

20-40 pi droplets on the parafilm and irradiated at a fluence of 15 W/m2 using germicidal lamps

(G15WT8, Sylvania) emitting predominantly at 254 nm. UV fluence was measured using a UVX

radiometer (UVP, San Gabriel, CA) with a 254 nm photocell (model UVX-25, UVP, San Gabriel,

CA).

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4. Materials and Methods

4.10. Photoreactivation

All experiments were done in a room equipped with yellow safety light (GE 'gold'

fluorescent light, Sylvania). An irradiation box equipped with 6 fluorescent lamps (Sylvania, 15

W F15T8 BLB, peak emission at 375 nm) and a metal rack connected to a tempered water bath

was used for photoreactivation. To keep the temperature of the samples around 30°C during

irradiation, the metal rack (17 cm away from the lamps) was set on 25-20°C. Irradiated

nucleosomes or naked DNA isolated from irradiated nucleosomes were mixed with E. coli

photolyase (Becton Dickinson) to yield a ratio of 70-75 ng photolyase/pg DNA. The

concentration ranges were 10-40 ng/pl DNA and 0.75-2 ng/pl photolyase in volumes of 80-100

pi. The mixture was pipetted into an inverted eppendorf tube cap, covered with a microscopic

slide, incubated at room temperature under yellow safety light for 5 minutes and irradiated at

fluences of 17-24 Wim2. The fluence was measured using a UVX radiometer (UVP, San Gabriel,

CA) with a 365 nm photocell (model UVX-36, UVP, San Gabriel, CA). To measure the

temperature during photoreactivation, an additional tube cap was filled with the same volume of

water and placed next to the others. The water temperature, measured with a needle thermometer,

was about 30°C during the irradiation. After various repair times, aliquots were taken for

nucleoprotein gel analysis (1.5-3 pi) and for mapping of the remaining CPDs by T4-endoV

cleavage (12-22 pi). The later samples were immediately mixed with 1.2 pi 10% SDS and kept

on ice in the dark until DNA isolation.

4.11. CPD Analysis

UV-irradiated and repair samples were adjusted to 0.5% SDS and Proteinase K (Roche)

was added to yield a concentration of 0.5 mg/ml. The samples were incubated for 3 hours at

50°C. The DNA was purified over Qiaquick PCR purification columns (QIAGEN) and eluted

with 50 pi 50 mM TRIS, 5 mM EDTA (pH 7.5). The samples were incubated for 5 minutes at

37°C, T4-endoV (Epicentre) was added (0.06 U T4-endoV/ng DNA) and incubation continued

for 2h at 37°C. 0.5-1 pi StrataClean Resin (STRATAGENE) was added to the samples. The

samples were vortexed and the volume adjusted to 100 pi with H20. After centrifugation in a

table top centrifuge at 13 000 rpm for 2 minutes, the supernatant was recovered and transferred

to a tube containing 10 pi NaAc pH 5.2. The DNA was precipitated by adding 400 pi ethanol,

vortexing, incubating the samples at -20°C for 1 hour and centrifiiging at 13 000 rpm and 4°C for

another hour. The pellets were washed once with 100 pi cold 70% ethanol and dried in the speed-

vac for 10 minutes. Scintillation counter (model 1500 Tri-Carb, Camberra Packard) was used to

103

4. Materials and Methods

determine the counts contained in each pellet, which were then solved in loading buffer (80%

formamide, 10 mM NaOH, 1 mM EDTA, 0.1% Xylene Cyanole, 0.1% Bromophenol blue) to

have the same counts/pl in every samples of one experiment (typically 2-7 x 103 cpm/pl). The

samples were analysed on 8% denaturing acrylamide gels.

4.12. Quantification of Nucleoprotein and Sequencing Gels

Dried gels were exposed to Phosphorimager storage screens (Molecular Dynamics) for 1

to 24 hours. The screens were scanned in a Phosphorlmager (Molecular Dynamics, model storm

820) at a resolution of 100 pm. All quantifications were done by volume integration using the

original Phosphorlmager files and the ImageQuant program (Molecular Dynamics, IQmac

version 1.2). The quantifications were done as previously described (Schieferstein, 1997).

Nucleoprotein gels were quantified as follows. Volume boxes were laid around the naked

DNA and nucleosomes bands. After subtraction of the background measured in an empty lane,

the ratio of nucleosomal bands to the sum of the nucleosomal and the naked DNA bands was

defined as reconstitution efficiency.

Sequencing gels of photoreactivation samples were quantified as follows. On the short-run

gels, volume boxes were laid around each CPD clusters (numbered from 1 to 19), the top band

and the entire lane. Gel background was measured in an empty lane and subtracted. The value of

each CPD cluster and top band was normalised to the signal of the entire lane to minimise the

influence of loading errors. Unspecific nicking background was obtained by laying volume

boxes corresponding the each CPD cluster and to the top band in the minus T4-endoV lane and

subtracted. The sum of all CPD cluster bands and of the top band was defined as 100% and the

percentage of CPDs measured in each cluster was calculated.

4.13. Gel Electrophoresis

4.13.1.LOW Melting Agarose Gels

0.8% - 1% low melting agarose (SeaPlaque agarose, FMC BioProducts) gels were run in

0.5 x TBE (45 mM Tris, 45 mM boric acid, 10 mM EDTA pH 8.0) at 4°C. DNA samples were

adjusted to 0.01% Bromophenol blue, 0.01% Xylene Cyanole, 6% glycerol, 0.5 x TBE prior to

loading.

104

4. Materials and Methods

4.13.2.Nucleoprotein Gels

Reconstituted nucleosomes were analysed on 4% acrylamide (19:1 acrylamide to Bis,

Appligene) gels in 0.5 x TBE (45 mM Tris, 45 mM boric acid, 10 mM EDTA pH 8.0). Where

indicated, the nucleoprotein gels also contained 10% glycerol (see figure legends). TE pH 7.5, 50

mM NaCl and 1 pi 5 x loading buffer (30% glycerol, 2.5 x TBE) were added to 0.5-2 pi of

reconstituted samples to yield a final volume of 5 pi and electrophoresed at 4°C for 3-4 hours at

10 mA constant current. The gels were dried on DE81 chromatographic paper (Whatman) for 1.5

hours at 80°C on a gel dryer (Biorad, model 583). The dried gels were exposed to X-ray films

(Fuji) and to phosporlmager plates (Molecular Dynamics).

4.13.3.Sequencing Gels

Cerenkov counting of the dried samples was done using a liquid scintillation analyzer

(model 1500 Tri-Carb, Canberra Packard). Samples were solved in sequencing loading buffer

(80% formamide, 10 mM NaOH, 1 mM EDTA, 0.1% Xylene Cyanole, 0.1% Bromophenol blue)

to a final concentration of 2-7 x 103 cpm/pl. The samples were denatured by incubation at 90°C

for 2 minutes and immediately chilled on ice. Samples (2 pl/lane) were analyzed on 8% (w/v)

Polyacrylamide (19:1 acrylamide to Bis, Appligene) gels (40 cm x 30 cm x 0.3 cm, BRL model

S2) containing 42% (w/v) urea in 1 x TBE (90 mM Tris, 90 mM boric acid, 20 mM EDTA pH

8.0). Gels were pre-run at 75 W (Biorad, model 3000 Xi) for 1-2 hours. Samples were loaded and

electrophoresed at 75 W (about 1800-2000 V, 40 mA) for the appropriate time period (ATDED:

short run: lh-2h, long run: 2h30-3h; ATDED-c: short run: Ihl5-lh30, long run: 2h30-3h;

ATDED-short: short run: lh-lhl5, long run: lh45-2h). The gels were dried on DE81 and 3MM

chromatographic paper (Whatman) for 2 hours at 80°C on a gel dryer (Biorad, model 583). The

dried gels were exposed to X-ray films (Fuji) and to phosporlmager plates (Molecular

Dynamics).

4.13.4.SDS-PAGE Gels

Histone proteins were analysed by SDS-PAGE using a 5% stacking gel and a 18%

resolving gel (29:1 acrylamide to Bis, ICN). Buffer concentration were as following: stacking gel

was 0.125 M TRIS, 1% SDS, pH 6.8; resolving gel buffer was 0.75 M TRIS, 1% SDS, pH 8.8;

electrophoresis buffer was 0.01 M TRIS, 0.08 M glycin, 0.1% SDS. SDS-sample buffer was

added to the proteins to yield final concentrations of 62 mM TRIS, 2.5% SDS, 9% glycerol, 0.35

M ß-mercaptoethanol, 0.001% Bromophenol blue, pH 6.8. The samples were boiled for 2

105

4. Materials and Methods

minutes at 95°C prior to loading. Samples were electrophoresed at 100 V for 2 hours. Gels were

stained in 10% TCA, 25% isopropanol, 0.1% Serva Blue R at RT over night and destained in

10% acetic acid, 25% methanol for 24 hours.

106

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125

Curriculum Vitae

Curriculum Vitae

Name

Present Adress

Date of Birth

Nationality

Current Position

Hélène Gaillard

Institute of Cell Biology, ETH-Hönggerberg, CH-8093 Zürich

Tel.: +41 1 633 33 43; Fax : +41 1 633 10 69

e-mail: helene.gaillard@cell.biol.ethz.ch

10th June 1975 in Lausanne

Swiss

Ph.D. Student

University Education

October 98 - April 02

November 93 - March 98

November 97 - February 98

September 96 - June 97

Ph.D. at the Institute of Cell Biology, ETH Zürich, in the

research group of Prof. Dr. Fritz Thoma.

Title: 'Nucleosome Remodeling Activities Act on UV-Damaged

Nucleosomes and Facilitate DNA-Repair'.

Diploma in Biology at the Biocentre, University of Basel.

Main subjects: Biochemistry, Biophysics, Cell Biology and

Microbiology.

Practical Training at the Department of Central Nervous System

Research, Hoffmann-La-Roche, Basel, in the research group of

Dr. Parisher Malherbe.

Diploma work at the Department of Microbiology, Biocentre,

University of Basel, in the research group of Prof. Dr. Charles

Thompson.

Title: 'Cloning and Characterisation of Regulatory Proteins

Involved in the Antibiotic Production of Streptomyces Species'.

School Education

August 90 - July 93

August 85 - July 90

'Matura Typus C (equivalent to A-Levels, with focus on

Sciences) at the Gymnase Cantonal de la Cité, Lausanne.

Secondary education at the Collège de Béthusy, Lausanne.

126

Curriculum Vitae

August 81 - July 85 Primary education at the Ecole Primaire de la Sallaz, Lausanne.

Fellowships

November 01-March 02 Roche Research Foundation

November 94-April 98 Bourse d'étude du Canton de Vaud

Language skills

French, German, English, Spanish

Oral or Poster Presentations

- Chromatin and Transcription Meeting, Snowmass, Colorado, USA, 2001

- RRR, Recombination, Replication and Repair Meeting, Zürich, 2001

- Swiss Workshop on genetic Recombination and DNA Repair, Les Diablerets, 2000

- ELSO, Geneva, 2000

- Symposium of the Departement of Biology, ETHZ, Davos, 2000

Scientific Publications

Gaillard, H., Fitzgerald, D., Smith, C. L., Peterson, C. L., Richmond, T. J., and Thoma, F.

Nucleosome remodeling activities act on UV-damaged nucleosomes and facilitate DNA-repair.

Submitted.

Folcher, M., Gaillard, H., Nguyen, L. T., Nguyen, K. T., LaCroix, P., Bamas-Jacques, N., Rinkel,

M., and Thompson, C. J. (2001). Pleiotropic functions of a Streptomyces pristinaespiralis

autoregulator receptor in development, antibiotic biosynthesis, and expression of a superoxide

dismutase. J Biol Chem 13, 13.

Malherbe, P., Richards, J. G., Gaillard, H., Thompson, A., Diener, C, Schüler, A., and Huber, G.

(1999). cDNA cloning of a novel secreted isoform of the human receptor for advanced glycation

end products and characterization of cells co- expressing cell-surface scavenger receptors and

Swedish mutant amyloid precursor protein. Brain Res Mol Brain Res 71, 159-170.

127