Structural and Functional Consequences of Platinum Anticancer Drug Binding to Free and
Nucleosomal DNA
by
MASSACHUSETS INSTITEf
OF TECHNOLOGY
Ryan Christopher Todd
JUN 0 2 2010
B.A. Chemistry (2003)
Johns Hopkins University
LIBRARIES
Submitted to the Department of Chemistry
in Partial Fulfillment of the Requirements for the Degree of
Doctor of Philosophy in Biological Chemistry
ARCHIVES
at the
Massachusetts Institute of Technology
June 2010
0 2010 Massachusetts Institute of Technology
All rights reserved
Signature of Author:
fRyan C. Todd
Department of Chemistry
May 10, 2010
Certified by:
'
'
- MW.pr
Stiphen(ILippard
Arthur Amos Noyes Professor of Chemistry
Thesis Supervisor
Accepted by:_
Robert W. Field
Haslam and Dewey Professor of Chemistry
Chairman, Departmental Committee on Graduate Studies
This doctoral thesis has been examined by a committee of the Department of Chemistry as
follows:
John M. Essigmann
William R. and Betsy P. Leitch Professor of Chemistry and Biological Engineering
Committee Chairman
I
-X
0
Okephen J. Lippard
Arthur Amos Noyes Professor of Chemistry
Thesis Supervisor
JoAnne Stubbe
Novartis Professor of Chemistry/Professor of Biology
Structural and functional consequences of platinum anticancer drug binding to free and
nucleosomal DNA
by
Ryan Christopher Todd
Submitted to the Department of Chemistry on May 10, 2010
in Partial Fulfillment of the Requirements for the Degree of
Doctor of Philosophy in Biological Chemistry at the
Massachusetts Institute of Technology
ABSTRACT
Cisplatin, carboplatin, and oxaliplatin are three FDA-approved members of the platinum
anticancer drug family. These compounds induce apoptosis in tumor cells by binding to nuclear
DNA, forming a variety of adducts, and triggering cellular responses, one of which is the
inhibition of transcription. The focus of this thesis is on studying the structure of these adducts,
and correlating these effects with inhibition of transcription. Chapter 1 presents (i) a detailed
review of the structural investigations of various Pt-DNA adducts and the effects of these lesions
on global DNA geometry; (ii) research detailing inhibition of cellular transcription by Pt-DNA
adducts; and (iii) a mechanistic analysis of how DNA structural distortions induced by platinum
damage may inhibit RNA synthesis in vivo. These hypotheses will be explored in subsequent
chapters of the thesis.
In Chapter 2, features of the 2.17 A resolution X-ray crystal structure of cisdiammine(pyridine)chloroplatinum(II) (cDPCP) bound in a monofunctional manner to
deoxyguanosine in a DNA duplex are discussed and compared to those of a cisplatin-1,2d(GpG) intrastrand cross-link in double-stranded DNA. The global geometry of cDPCPdamaged DNA is quite different from that of DNA containing a cisplatin 1,2-d(GpG) cross-link.
The latter platinated duplex is bent by ~40* toward the major groove at the site of the adduct;
however, the monofunctional Pt-dG lesion causes no significant bending of the double helix.
Like the cisplatin intrastrand adduct, however, the cDPCP moiety creates a distorted base pair
step to the 5' side of the platinum site that may be correlated to its ability to destroy cancer cells.
Structural features of monofunctional platinum adducts are analyzed, the results of which
suggest that such adducts may provide a new platform for the design and synthesis of Pt
anticancer drug candidates.
The role of carbonate in the binding of cis-diamminedichloroplatinum(II) to DNA was
investigated in Chapter 3 in order to understand the potential involvement of carbonato-cisplatin
species in the mechanism of action of platinum anticancer agents. Cisplatin was allowed to react
with single-stranded DNA in carbonate, phosphate, and HEPES buffers, and the products were
analyzed by enzymatic digestion/mass spectrometry. The data from these experiments
demonstrate (1) that carbonate, like other biological nucleophiles, forms relatively inert
complexes with platinum that inactivate cisplatin, and (2) that the major cisplatin-DNA adduct
formed is a bifunctional cross-link. These results are in accord with previous studies of cisplatinDNA binding and reveal that the presence of carbonate has no consequence on the nature of the
resulting adducts.
The 1.77-A resolution X-ray crystal structure of a dodecamer DNA duplex with the
sequence 5'-CCTCTGGTCTCC-3' that has been modified to contain a single engineered 1,2-cis{Pt(NH 3)2}2+-d(GpG) cross-link, the major DNA adduct of cisplatin, is reported in Chapter 4.
These data represent a significant improvement in resolution over the previously published 2.6-A
structure. The ammine ligands in this structure are clearly resolved, leading to improved
visualization of the cross-link geometry with respect to both the platinum center and to the
nucleobases, which adopt a higher energy conformation. Also better resolved are the deoxyribose
sugar puckers, which allow us to re-examine the global structure of platinum-modified DNA.
Another new feature of this model is the location of four octahedral [Mg(H 2 0) 6 ]2+ ion associated
with bases in the DNA major groove and the identification of 124 ordered water molecules that
participate in hydrogen bonding interactions with either the nucleic acid or the
diammineplatinum(II) moiety.
Chapter 5 discusses structural investigations of nucleosomal DNA modified by sitespecific platinum adducts. Nucleosome core particles containing a single 1,3-cis-{Pt(NH 3)2}2+_
d(GpTpG) intrastrand cross-link were synthesized and crystallized, and the X-ray structure was
determined at 3.2 A resolution. The cisplatin adduct adopts a conformation facing toward the
octamer core, in agreement with previous experiments. DNA in the vicinity of the Pt adduct has
a similar helical bend as that observed in the NMR solution structure of free DNA containing the
same cross-link, indicating that the rotational positioning power of cisplatin intrastrand crosslinks stems from the propensity to align the bent Pt-DNA structure with the DNA curvature
arising from the nucleosome superhelix.
Functional consequences of cisplatin binding to nucleosomal DNA are explored in
Chapter 6. The effect of a single engineered platinum intrastrand cross-link on ATPindependent nucleosome mobility was investigated in vitro. Both 1,2-d(GpG) and 1,3-d(GpTpG)
adducts of cisplatin inhibit translocation of DNA along the histone octamer, with the former Pt
lesion providing a larger barrier. In vitro transcription assays with T7 RNA polymerase were
conducted to determine whether cisplatin-DNA cross-links inhibit RNA synthesis by preventing
access to nucleosomal DNA. Immobilized transcription templates containing a T7 RNAP
promoter site, a single engineered cisplatin 1,2-d(GpG) or 1,3-d(GpTpG) intrastrand cross-link,
and a phased nucleosome core particle were prepared. Analysis of resulting RNA transcript
length revealed that the T7 RNAP elongation complex can overcome the energy barrier to
nucleosome sliding caused by platinum intrastrand cross-links, but stalls when it reaches a PtDNA adduct placed on the DNA template strand. These results provide further evidence that
intrastrand cross-links of cisplatin inhibit transcription by creating a physical barrier that the
polymerase cannot pass.
Appendices A and B summarize incomplete work that may be of use to future
researchers working in this area. Appendix A describes attempts to isolate isomerically pure PtDNA probes containing a photoreactive benzophenone moiety, for use in cross-linking
experiments that identify proteins that recognize and interact with cisplatin-DNA damage. In
Appendix B efforts to obtain an X-ray crystal structure of an 1 Imer duplex DNA containing the
1,3- cis-{Pt(NH 3)2 }2 -d(GpTpG) intrastrand cross-link are reported. Appendix C details HPLC
and mass spectrometric methods for purification and analysis of platinated oligonucleotides that
were developed in the course of this research.
Thesis Supervisor: Stephen J. Lippard
Title: Arthur Amos Noyes Professor of Chemistry
ACKNOWLEDGEMENTS
Graduate school at MIT is often characterized as a lengthy and grueling ordeal, where
those fortunate enough to complete their program wear the letters "PhD" like a badge of honor,
having survived a horrific initiation into the club. For me graduate school was every bit as
difficult as I expected it to be; there were times when I felt as though I would never complete my
degree. However, even in the worst periods of frustration, life as a graduate student was not as
painful as others describe their experiences. Many individuals deserve credit for helping me
through the last five years at MIT, and for making graduate school more of a fruitful learning
experience and less of an excruciating marathon.
I am exceptionally grateful to my advisor Stephen J. Lippard for the support and
guidance I've received throughout my graduate career. As a scientist Steve runs a diverse
laboratory that is sufficiently equipped for either synthetic inorganic research or cell biology
work. The variety of science to which I've been exposed over the last five years has significantly
increased my breadth as a chemist. Nearly every experiment in this thesis, from gel
electrophoresis of DNA to synthesis of platinum coordination compounds to macromolecular Xray crystallography, was something I learned and performed for the first time in Steve's
laboratory. As an advisor, Steve achieved a nearly perfect balance between being accessible to
provide regular guidance on research plans and advice on troubleshooting experiments, and
allowing me the freedom to work through issues, make mistakes, and learn on my own. After
leaving MIT I probably will not continue working in X-ray crystallography or even platinum
anticancer research, but the knowledge that I gained about these fields was insignificant relative
to the general scientific training I received under Steve's tutelage. Critical analysis of data and
clear communication of results are universal skills that are absolutely necessary for any field of
laboratory science, and these are the most valuable abilities that I take away from the Lippard
lab.
In addition to Steve, many other members of the laboratory provided help and support
along the way. Evan Guggenheim first helped get me settled into the lab and guided me through
many of my initial experiments. Mike McCormick served as my initial mentor in X-ray
crystallography. In addition, these two were also good friends in the group that provided valuable
advice in navigating through year-end reports, group meetings, and other responsibilities
associated with Steve's group, as well as stress relief in the form of lunches and happy hours.
Christy Tinberg and Lindsey McQuade, fellow members of the Lippard Lab Class of 2010,
provided moral support and scientific feedback throughout classes, cumes, orals exams, and our
other graduate school milestones. I wish them the best of luck in their respective post-doctoral
endeavors. Myriad members of the lab and the platinum subgroup have helped me through the
years in troubleshooting experiments, designing research plans, editing papers, and interpreting
results. These individuals include Matthias Ober, Datong Song, Wee Han Ang, and Guangyu
Zhu, among others. Finally, I need to acknowledge my research technician Paresh Agarwal, who
helped to execute some of the experiments documented in this thesis. In his year of service in the
Lippard lab between graduation from MIT and entering graduate school at UC-Berkeley, he was
an excellent labmate who picked up skills quickly and performed experiments carefully and
thoughtfully.
The Lippard lab is not a specialized unit for macromolecular X-ray crystallography, as
are most groups that conduct research in this field; we simply utilize it as one of the many
possible tools for investigating a biochemical system. Thus, collaboration was a necessary
component of my research. I worked closely with several experts in the field in the course of my
work. Alejandro D'Aquino, a postdoctoral associate in the laboratory of Greg Petsko at Brandeis
University, taught me everything about phasing a crystal structure by anomalous signal, as well
as structure refinement in general, and played an integral role in completing and publishing my
first DNA structure. Kanagalaghatta Rajashankar and Narayanasami Sukumar at the NE-CAT
beamline at the Advanced Photon Source, Argonne National Laboratory, and Tzanko Doukov at
the Stanford Synchrotron Radiation Laboratory are beamline scientists who aided in diffraction
data collection at these sites. Members of Cathy Drennan's lab in the chemistry department,
including Leah Blasiak, Ainsley Davis, and Yan Kung, are also acknowledged for helpful
discussions.
I am grateful to members of the MIT chemistry department for their roles in supporting
me through the program. My committee chairman John Essigmann has provided valuable and
insightful scientific feedback during our annual meetings, in addition to personal and
professional guidance. Professor JoAnne Stubbe, in addition to being one my of committee
members, was an excellent teacher who, between 5.50, Enzymes: Structure and Function, and
5.52, Advanced Biological Chemistry, taught me more about biological chemistry in one
semester than I had yet learned in my life. I also thank Susan Brighton, Chemistry Graduate
Administrator, for all of her help in various events throughout school.
Many other scientists served as mentors to me earlier in my research career. Robert
Johnston took me into an analytical laboratory in high school and introduced me to FTIR, GC,
and column chromatography, as well as providing a source of income so that I could work
summers during college. David Goldberg, my undergraduate advisor at Johns Hopkins, provided
me the first opportunity to work in a research lab. My supervisor at Merck & Co., Inc., Randy
Seburg, taught me many principles of experimental design, method development, and
documentation of research, without which I would have struggled though graduate school.
As evidenced by the list above, many people played a role in my scientific development
over the last five years. Completion of my thesis would not have been possible without the
excellent chemists around me every day. However, equal credit for this accomplishment has to
be given to the personal support I've received from my friends and family. Members of Twisted
Metal and Ironside Ultimate served as friends and teammates, and provided an outlet for me to
spend time and energy away from the lab, which helped me to remain focused and engaged
while at work. All of my family members, including my parents, brothers and sisters, and inlaws, have been fully encouraging of my endeavors, and I am grateful for their support.
Finally, I need to thank my lovely wife Moira Todd, who more than anyone, supported
me during every day of my graduate career. She dealt with many dinners alone while I was in
lab, and kept our house clean and running smoothly while I was busy with work. She calmed me
when I was frustrated, and celebrated with me when milestones were completed or papers
published. As a major in international relations, she did not understand most of my research, but
that never stopped her from asking about my day. She always remained involved and interested
in my work. She even supported me financially so that we could live above the means of my
lowly grad school stipend. I now look forward to moving on to the next phase of our lives
together. Thanks Moira, I love you.
TABLE OF CONTENTS
ABSTRACT
3
ACKNOWLEDGEMENTS
6
TABLE OF CONTENTS
8
LIST OF TABLES
14
LIST OF SCHEMES
15
LIST OF FIGURES
16
Chapter 1: Inhibition of Transcription by Platinum Antitumor Compounds
20
INTRODUCTION
22
DNA ADDUCTS FORMED BY PLATINUM ANTITUMOR AGENTS
Cisplatin/Carboplatin
Oxaliplatin
Non-classical platinum compounds
Effects of platinum binding on nucleosome structure
24
24
27
29
32
PROTEIN BINDING TO PLATINUM-DNA ADDUCTS
Upstream binding factor
TATA-binding protein
Y-box binding protein
High-mobility group box protein 1
Stucture specific recognition protein 1
Poly(ADP-ribose) polymerase I
34
34
35
35
35
36
37
INHIBITION OF TRANSCRIPTION BY PLATINUM ANTITUMOR COMPLEXES
Reconstituted systems
Studies in cell extracts and cell culture
Transcription-coupled repair of Pt-DNA adducts
37
38
39
41
THE MECHANISM OF TRANSCRIPTION INHIBITION BY PT-DNA ADDUCTS
Transcription factor hijacking
Roadblock of RNA polymerases
Disruption of chromatin dynamics
43
43
44
45
FROM TRANSCRIPTION INHIBITION TO APOPTOSIS
47
FUTURE DIRECTIONS
48
REFERENCES
Chapter 2. X-Ray Crystal Structure of a Monofunctional Platinum-DNA Adduct,
cis-{Pt(NH 3 )2(pyridine)}2+Bound to Deoxyguanosine in a Dodecamer Duplex
61
INTRODUCTION
62
EXPERIMENTAL
Materials
Synthesis of cis-diammine(pyridine)chloroplatinum(II)
Deoxyoligonucleotide synthesis and purification
Synthesis of G6-platinated oligonucleotide
Synthesis of G7-platinated oligonucleotide
Preparation of Pt-DNA duplexes
Crystallization and X-ray data collection
Structure determination and refinement
64
64
64
66
66
68
68
69
70
RESULTS
Sample preparation and crystallization
Unit cell composition and crystal packing
Platinated DNA duplex
72
72
73
74
DISCUSSION
77
CONCLUSIONS
82
REFERENCES
83
Chapter 3. Reactions Between Phosphate and Carbonate Complexes of Cisplatin
and Nucleic Acids: Investigations of Resulting Pt-DNA Adduct Structure and Yield
86
INTRODUCTION
87
EXPERIMENTAL
Materials
Reaction of cisplatin with single-stranded DNA
Mass spectrometry
Enzymatic digestion
88
88
88
89
89
RESULTS
Product characterization
Yield of Pt-DNA adducts
90
90
93
DISCUSSION
95
CONCLUSIONS
REFERENCES
100
Chapter 4. Structure of Duplex DNA Containing the Cisplatin 1,2-{Pt(NH 3)2} 2+_
d(GpG) Cross-Link at 1.77 A Resolution
102
INTRODUCTION
103
EXPERIMENTAL
Materials
Preparation of platinated DNA duplex
Crystallization and X-ray diffraction data collection
Structure determination and evaluation
104
104
105
106
107
RESULTS AND DISCUSSION
Unit cell and crystal packing
Global DNA geometry
Pt adduct geometry
Magnesium site identified
Ordered water molecules
109
109
110
113
116
117
CONCLUSIONS
119
REFERENCES
121
Chapter 5. Structural Investigations of a Site-Specifically Platinated Nucleosome
Core Particle Containing the Cisplatin 1,3-{Pt(NH 3 )2}2+-d(GpTpG) Cross-Link
131
INTRODUCTION
132
EXPERIMENTAL
Materials
Oligonucleotide synthesis
Characterization of wl-Pt and w2-Pt: MALDI-TOF mass spectrometry
Characterization of wl-Pt and w2-Pt: Nuclease SI/CIP digestion
Pt-DNA adduct stability test
Synthesis of site-specifically platinated DNA duplexes tI-Pt and t2-Pt
Restriction enzyme digestion of tl-Pt/t2-Pt
Purification of histone core proteins H2A, H2B
Histone octamer refolding/purification
Assembly of nucleosome core particles
Crystallization studies
Data collection and processing
Model refinement
136
136
137
139
139
140
140
141
141
143
143
145
146
148
RESULTS
Synthesis of site-specifically platinated mononucleosomes
Crystallization and data collection
Structure of the platinated nucleosome
149
149
151
152
DISCUSSION
157
CONCLUSIONS
160
REFERENCES
162
Chapter 6. Exploring Transcription by T7 RNA Polymerase of Free and
Nucleosomal DNA Modified with Site-Specific Platinum IntrastrandCross-Links
165
INTRODUCTION
166
EXPERIMENTAL
Materials
Preparation of s1/s2/s 1-Pt/s2-Pt duplexes
Preparation of c l/c2/c 1-Pt/c2-Pt duplexes
Assembly and purification of nucleosome core particles
Nucleosome mobility investigation
Preparation of 204 bp immobilized transcription templates
Restriction enzyme mapping of nucleosome position
Single-round in vitro transcription assays with T7 RNA polymerase
169
169
170
172
174
175
175
177
178
RESULTS AND DISCUSSION
Preparation of immobilized, site-specifically platinated free and nucleosomal DNA
transcription templates
Nucleosome mobility investigation
Single-round in vitro transcription assays
180
180
183
186
CONCLUSION
191
REFERENCES
193
RESEARCH SUMMARY AND PERSPECTIVES
195
Appendix A. Towards Separation of PtBP6-DNA Orientational Isomers by HPLC
197
INTRODUCTION
198
EXPERIMENTAL
Materials
Platination of 14mer with PtBP6
Resolution of orientational isomers by HPLC
199
199
200
200
RESULTS AND DISCUSSIONS
203
CONCLUSIONS
204
REFERENCES
205
Appendix B. Crystallization Attempts of a DNA 11mer Duplex Containing a SiteSpecific 1,3-cis-{Pt(NH 3)2}2+-d(GpTpG) Lesion
206
INTRODUCTION
207
EXPERIMENTAL
Materials
Synthesis and purification of 1 imer duplex containing a 1,3-cis-{Pt(NH 3)2}2 +_
d(GpTpG) cross-link
Crystallization attempts of Pt-1 1mer duplex
208
208
RESULTS AND DISCUSSION
Synthesis/purification of IlImer duplex w/ 1,3-cis-{Pt(NH 3)2}2+-d(GTG) adduct
Crystallization attempts of Pt-1 1mer duplex
211
211
212
CONCLUSION
212
REFERENCES
213
Appendix C: HPLC Methods for Purification and Analysis of Platinated
Oligonucleotides
215
RCT.001P: Purification of dimethoxytrityl-containing short oligonucleotides from untritylated failure sequences by preparative reverse-phase HPLC
RCT.002S: Purification of 12mer DNA platination reactions by semi-preparative ionexchange HPLC
RCT.003S: Purification of dodecamer duplex DNA containing a single platinum-DNA
adduct by semi-preparative ion-exchange HPLC
RCT.004A: Analysis and purification of 14mer DNA platination reactions with cisplatin
by ion-exchange HPLC
RCT.005S: Purification of dimethoxytrityl-containing short oligonucleotides from untritylated failure sequences by semi-preparative reverse-phase HPLC
RCT.006S: Purification of 14mer DNA platination reactions by semi-preparative ionexchange HPLC
RCT.007A: Purification of 14mer DNA platination reactions by analytical ion-exchange
HPLC
RCT.008A: Analysis of platinated oligonucleotides digested with nuclease Sl/Pl and
calf intestinal phosphatase by reverse-phase HPLC
209
209
216
217
218
219
220
221
222
223
RCT.009M: Analysis of platinated oligonucleotides digested with nuclease SI/PI and
calf intestinal phosphatase by reverse-phase HPLC/ESI-MS
224
RCT.010M: Electrospray mass spectrometry of oligonucleotides with on-line HPLC
desalting
226
Biography
227
Curriculum Vitae
228
LIST OF TABLES
Table 1.1. Structural features of platinum-DNA adducts.
33
Table 2.1. Collection and refinement statistics for the pyriplatin-DNA crystal structure
72
Table 2.2. Geometric parameters for pyriplatin- and cisplatin-DNA duplexes
76
Table 4.1. Collection and refinement statistics for the cisplatin-DNA crystal structure
108
Table 4.2. Geometry comparisons of the 1,2-cis-{Pt(NH 3)2}2 -d(GpG) cross-link
114
Table 4.JS. List of hydrogen-bonding interactions in the cisplatin-DNA crystal structure
125
Table 5.1. Data collection statistics for tl-Pt-NCP crystals frozen under different
cryoconditions.
147
Table 5.2. Comparison of diffraction data sets truncated spherically at 3.9 A, or
ellipsoidally between 3.2 and 4.1 A.
148
Table 5.3. Refinement statistics for tl-Pt-NCP structure.
149
Table 6.1. Restriction enzyme digestion of 204-bp transcription templates
182
LIST OF SCHEMES
Scheme 2.1. Synthesis of cis-diammine(pyridine)chloroplatinum(II) from K2 PtCl 4.
LIST OF FIGURES
Figure 1.1. Chemical structures of platinum anticancer agents
22
Figure 1.2. X-ray crystal and NMR structures of double stranded DNA containing
adducts of various platinum anticancer agents
28
Figure 1.3. Protein recognition and binding to Pt-DNA adducts
36
Figure 1.4. Possible mechanisms of transcription inhibition by platinum antitumor
agents
44
Figure 2.1. Chemical structures of cisplatin, oxaliplatin, and pyriplatin
63
Figure 2.2. HPLC chromatograms of the purification of single- and double-stranded
platinated dodecamer DNA
67
Figure 2.3. ESI-MS spectra of DNA strands
68
Figure 2.4. HPLC traces of enzymatic digestion analysis with nuclease P1 and calf
intestinal phosphatase
69
Figure 2.5. X-ray diffraction image of crystals of a pyriplatin-DNA duplex, and solventflattened electron density map
71
Figure 2.6. Crystal packing interactions between pyriplatin-DNA molecules
74
Figure 2.7. The structures of (a) cDPCP- and (b) cDDP-damaged DNA duplexes
75
Figure 2.8. Stereoscopic views of the cDPCP-dG adduct on duplex DNA
76
Figure 2.9. The pyriplatin-DNA platinated base pair
78
Figure 2.10. Active sites of RNA polymerase II stalled at a cDPCP-dG adduct
81
Figure 3.1. HPLC chromatograms of the reaction of cisplatin with 14mer DNA
90
Figure 3.2. ESI (-) spectrum of major product of cisplatin-DNA reaction
91
Figure 3.3. HPLC chromatograms of enzymatic digestion of 14mer DNA, the major
platinated product and the minor Pt speices
92
Fig. 3.4. ESI (-) mass spectra of Pt-DNA cross-links arising from enzymatic digestion
of single-stranded DNA
93
Figure 3.5. Yield of platinated 14mer ssDNA in 24 mM buffer
94
Figure 3.6. Routes for biological processing of cisplatin
98
Figure 4.1. HPLC chromatograms of the purification of single- and double-stranded
platinated dodecamer DNA
106
Figure 4.2. Structural features of cisplatin-damaged DNA
110
Figure 4.3. Stereo view of the previously published structure of DNA modified with a
1,2-cis- {Pt(NH 3)2 }2+-d(GpG) cross-link superimposed on the current structure
111
Figure 4.4. Stereo images of 2F-Fe electron density maps defining deoxyribose sugar
conformations
112
Figure 4.5. Views of the 1,2-cis-{Pt(NH 3)2}2+-d(GpG) adduct
115
Figure 4.6. Binding of [Mg(H 2 0) 6 ]2 +cations to purine dinucleotides in the cisplatinDNA dodecamer duplex
117
Figure 4.7. Schematic depicting hydrogen-bonding interactions between solvent
molecules and Pt-DNA
118
Figure 4.3S. Base pair step parameters for cisplatin-DNA duplexes
124
Figure 5.1. Oligonucleotides synthesized towards ligation of 146 bp DNA containing
single cisplatin cross-links
138
Figure 5.2. HPLC purification of wl-Pt from side products
138
Figure 5.3. HPLC purification of w2-Pt from side products
139
Figure 5.4. SDS-PAGE analysis of fractions for histone octamer purification
143
Figure 5.5. Purification of t1-Pt-NCP by preparative gel electrophoresis
145
Figure 5.6. Diffraction image of t1-Pt-NCP crystal and signal-to-noise ratios (FAY) of
diffraction data in the a*, b*, and c* directions
148
Figure 5.7. Restriction enzyme digestion of ti, ti-Pt, t2, or t2-Pt.
151
Figure 5.8. Structure of the a platinum-damaged nucleosome core particle
Figure 5.9. Overlay of platinated nucleosomal DNA with DNA from the original
nucleosome structure
153
Figure 5.10. B-factor distribution of phosphorus atoms of the DNA backbone
154
154
Figure 5.11. Possible 1,3-cis-{Pt(NH 3)2}2+-d(GpTpG) cross-link locations on the
nucleosome core particle.
156
Figure 5.12. Stereo view of the 1,3-cis-{Pt(NH 3)2}2+-d(GpTpG) cross-link, looking
down the DNA double helix
157
Figure 5.13. Global helical bend angle of the NMR solution structure of duplex DNA
containing the l,3-cis-{Pt(NH 3)2 }2+-d(GTG) cross-link and a DNA segment containing
the adduct taken from the nucleosome structure
159
Figure 5.14. Analysis of roll angle of the DNA superhelix in the high-resolution X-ray
crystal structure of the nucleosome core particle
160
Figure 6.1. Mechanisms of nucleosomal transcription
167
Figure 6.2. Oligonucleotides synthesized towards ligation of nucleosomal DNA
containing single cisplatin cross-links on the template strand
171
Figure 6.3. HPLC purification of dl-Pt
172
Figure 6.4. HPLC purification of d2-Pt
172
Figure 6.5. Oligonucleotides synthesized towards ligation of nucleosomal DNA
containing single cisplatin cross-links on the coding (non-template) strand
173
Figure 6.6. Assembly of nucleosome core particles of 145 bp DNA duplexes
175
Figure 6.7. Ligation of 204-bp transcription templates with free or nucleosomal DNA
177
Figure 6.8. Restriction enzyme sites along the 204-bp DNA
178
Figure 6.9. Experimental system using immobilized free or nucleosomal templates to
study transcription by T7 RNA polymerase
179
Figure 6.10. Gel analysis of restriction enzyme mapping of nucleosome core particle
position
183
Figure 6.11. Native PAGE analysis of nucleosome mobility investigation of platinated
nucleosome core particles
184
Figure 6.12. Quantitation of nucleosome mobility of platinated samples at either 37*C
or 50 *C
185
Figure 6.13. Transcription by T7 RNA polymerase of 204-bp templates containing free
or nucleosomal DNA containing no platinum adduct, a 1,3-cis-Pt(GTG) cross-link, or a
1,2-cis-Pt(GG) cross-link on either the template or coding strand
187
Figure 6.14. Kinetics of transcription by T7 RNA polymerase of 204-bp templates
containing free or nucleosomal DNA containing no platinum adduct, a 1,3-cis-Pt(GTG)
cross-link, or a 1,2-cis-Pt(GG) cross-link on the template strand
188
Figure 6.15. Inhibition of transcription by T7 RNA polymerase from site-specific 1,3cis- {Pt(NH 3)2 }2+-d(GpTpG) or 1,2-cis-{Pt(NH 3)2}2+-d(GpG) cross-links
188
Figure A.1. The structure of PtBP6, and depiction of the two orientational isomers of
PtBP6 on 14mer single-stranded DNA
199
Figure A.2. HPLC method and chromatogram for purification of PtBP6-14mer from
unplatinated oligonucleotide using a C4 reverse-phase column
201
Figure A.3. HPLC method and chromatogram for purification of PtBP6-14mer from
unplatinated oligonucleotide using a C18 reverse-phase column and 50 mM TEAA
202
Figure A.4. HPLC method and chromatogram for purification of PtBP6-14mer from
unplatinated oligonucleotide using a C18 reverse-phase column and 100 mM TEAA
203
Figure B.1. Stereo view of the NMR solution structure of duplex DNA containing a
cisplatin 1,3-cis-{Pt(NH3)2 }2+-d(GTG) intrastrand cross-link
207
Figure B.2. Purification of single- and double-stranded 11Imer platinated DNA
210
Chapter 1. Inhibition of Transcription by Platinum Antitumor Compounds
Portions of this chapter were published as a critical review in Metallomics, 2009, 1 (4) 280-291.
Metallomics
Lww~ nrItiwa
flamkcr
Vnl2
NL2Y(
CRITICAL REVIEW
RSC
CRIT1ICAL
REVIEW11111l
tid
"V
-
MtF tAf
-q
-z~
.
-
JLV,:3
Paotf1-ir
iIII
Cover image of Metallomics, 2009, 1 (4) featuring this review of transcription inhibition by
platinum antitumor drugs.
Introduction
One of the great success stories in the field of cancer chemotherapy is that of cisplatin
(cis-diamminedichloroplatinum(II), or cis-DDP), a curative treatment for testicular tumors.' Approved by the U.S. Food and Drug Administration in 1978, cisplatin is also administered for several other forms of cancer, including ovarian, cervical, head and neck, esophageal, and nonsmall-cell lung cancers. 1-3 Only in testicular cancer, however, does the drug reach greater than
Treatment can be limited by toxic side
90% cure rates, approaching 100% in early stage cases.'
effects, including nephrotoxicity, emetogenesis, and neurotoxicity.' Resistance to the drug, either
acquired or inherent, is also common. Two other members of the platinum antitumor drug family, carboplatin and oxaliplatin (Fig. 1.1), have subsequently been approved for use in the United
States. Whereas carboplatin and cisplatin are cross-resistant,7 oxaliplatin has a different spectrum
of activity and has become a first-line therapy for colorectal cancer.
0
HSN/
HN
Pt
CI
\C
HN
Pt
H3N'
H2
ON
Pt
"'
O0
O
0
H2
Oxaliplatin
Carboplatin
Claplatin
O
0
OK+
WN I "CI
N't
H2 0
7
NHa
CIN
CI
NH3
0
Satraplatin
Pyriplatin
Figure 1.1. Chemical structures of platinum anticancer agents. Cisplatin, carboplatin, and oxaliplatin are FDA-approved for chemotherapy use in the United States. Satraplatin was the first
Pt(IV) complex to reach Phase III clinical trials as an orally available platinum compound.
Pyriplatin is a non-classical, monofunctional platinum complex with antitumor activity in colorectal cancer cells that inhibits transcription in vitro.
Since the serendipitous discovery of its antineoplastic activity, 9"10 many research groups
have focused on revealing the molecular details of the mechanism of action of cisplatin and related compounds. The early steps of triggering cell death by platinum(II) compounds involve
four stages. They are (1) cellular accumulation by both passive and active uptake; (2) activation
of the platinum(II) complex; (3) binding to nucleic acids to form a variety of Pt-DNA adducts;
and (4) the cellular response to DNA damage."'"2 For years it was thought that cisplatin entered
cells primarily through passive diffusion, owing to data that showed platinum uptake was neither
saturable nor inhibited by structural analogues.13- 5 However, a growing body of evidence suggests a role for active uptake by membrane proteins, such as the copper transporter CTR1, in cisplatin accumulation.161 7 Cisplatin activation involves replacement of the chloride ligands with
water molecules in consecutive first order processes, driven by a drop in chloride ion concentration as the compound crosses the cell membrane.' 8 The aquated forms of cisplatin bind DNA at
the N7 position of purine bases to form primarily 1,2-intrastrand adducts between adjacent
guanosine residues.19 A smaller number of 1,3-intrastrand, interstrand, and monofunctional PtDNA adducts also form. The DNA damage leads to disruption of several cellular processes including transcription and replication. After cell cycle arrest, the Pt lesions are either removed by
nucleotide excision repair or apoptosis is triggered.
Early mechanistic studies led to the formation of several classical structure-activity relationships.2 0 In particular, it was hypothesized that an active platinum antitumor complex should
have square-planar geometry, contain two labile leaving groups in a cis conformation, be neutral
to facilitate passive diffusion across cell membranes, and contain inert amine ligands in the nonleaving-group positions. Since that time, however, many non-classical "rule-breakers," including
polynuclear platinum compounds, 2' platinum(IV) complexes,2 2 monofunctional platinum(II)
complexes,2 3 and compounds with trans sterochemistry,2 4 2 5 have been discovered with significant ability to destroy cancer cells.
The scope of this thesis is limited to the DNA binding and cellular processing aspects of
the mechanism of action of platinum anticancer complexes. In this chapter structural studies of
various DNA adducts that arise from binding different members of the platinum antitumor drug
family are reviewed. Cisplatin and related complexes bound to DNA have been thoroughly studied by both X-ray crystallography and NMR spectroscopy, yielding abundant information about
platinum modification of DNA structure. Next, studies demonstrating that cisplatin blocks transcription are described and data that implicate transcription inhibition as a major pathway involved in cancer cell death are discussed. Finally, the current hypotheses detailing how platinumDNA adducts block transcription by RNA polymerases are introduced, including how this disruption can promote apoptosis through p53-dependent and -independent pathways. These theories will be explored in subsequent chapters of the thesis.
DNA Adducts Formed by Platinum Antitumor Agents
Cisplatin/Carboplatin. Cisplatin forms a spectrum of intra- and interstrand DNA cross-links
which have been identified both in vitro and in vivo. 26-29 The major adduct, comprising -65% of
total products, is a 1,2- {Pt(NH 3)2 }2 -d(GpG) intrastrand cross-link. Other minor products include
1,2-d(ApG) (~25%) and 1,3-d(GpNpG) (5-10%) intrastrand adducts, as well as a smaller number
of interstrand cross-links (ICL) and monodentate adducts. Surprisingly, the 1,2-d(GpA) lesion is
not observed either in vitro or in vivo. 30 Although carboplatin forms the same type of adducts as
cisplatin, the product profile is markedly different in cells. 3 ' The major carboplatin adduct identified was cis-[Pt(NH 3)2 (dG) 2] (36%), which could arise from 1,3-d(GpNpG) intrastrand cross-
links. Minor products included 1,2-d(GpG) (30%), 1,2-d(ApG) (16%), as well as a small number
of interstrand (3-4%) cross-links and monofunctional adducts. Because trans-diammine32,33
and because
dichloroplatinum(II) is incapable of forming 1,2-intrastrand cross-links on DNA,3'3
this complex has insignificant antitumor activity in cells, 34 intrastrand adducts are more likely to
be responsible for the cytotoxicity of cisplatin in cancer cells. Further investigations of platinum
interstrand cross-links revealed no correlation between the frequency of ICL formation and cytotoxicity, providing additional evidence that intrastrand cross-links are essential to tumor cell
death."
The formation of cisplatin-DNA cross-links structurally distorts the DNA. Initial biochemical experiments showed that cisplatin binding unwinds DNA36-38 and results in a loss of
helix stability, as demonstrated by calorimetric studies 39-41 and denaturing gel electrophoresis
studies. 42 Further calorimetric experiments with site-specific cisplatin-DNA adducts revealed a
duplex destabilization of 6.3 kcal/mol associated with 1,2-d(GpG) adduct formation.4 3 The extent
of this destabilization was subsequently shown to be sequence dependent.4 4
Gel electrophoresis was also utilized to measure bending of DNA by various cisplatinDNA cross-links. 4 5-4 7 1,2-Intrastrand cross-links bend the helix by 32*-34* and unwind it by 130,
whereas 1,3-intrastrand adducts bend DNA by 350 and unwind it by 230. In similar studies per-
formed with interstrand cross-links formed by cisplatin binding to two guanines, the DNA was
bent by 45-55* toward the major groove and unwound by 790.48'49
X-ray structural investigations of Pt-DNA adducts initially focused on platinated di- or
trinucleotides.
50-52
However, it was not until a platinated DNA dodecamer duplex containing a
site-specific 1,2-{Pt(NH 3)2 }2+-d(GpG) intrastrand cross-link was solved by X-ray crystallography that fine details of the structure of Pt-damaged DNA began to emerge (see Fig. 1.2.a).53 '54
The X-ray crystal structure revealed that the Pt adduct induces a global bend in the DNA duplex
by 35-40* and unwinds the double helix by ~25'. The major groove is compacted and the minor
groove widened and flattened. The DNA takes on A-form properties to the 5' side of the Pt
cross-link and a B-form structure on the 3' side of the 1,2-d(GpG) adduct. The roll angle between platinated guanine bases is 260. This relatively shallow roll angle results in considerable
strain being placed on the Pt-N7 bonds, displacing the Pt atom out of the guanine ring planes by
approximately 1 A each. Subsequent NMR spectroscopic studies 55' 56 revealed differences between the solid state and solution structures, which could be traced to crystal packing interactions in the former. The solution structures showed bend angles of 60-70' and an exaggerated
roll of 490 at the 1,2-{Pt(NH 3)2}2+ d(GpG) cross-link. In addition, the NMR structures contained
primarily B-form DNA.
Examination of the NMR structures of duplex DNA containing a 1,2-{Pt(NH 3)2}2 +_
d(GpG) cross-link revealed significant distortion of the DNA base pair step to the 5' side of the
adduct.57' 58 This conformational change is marked by unusually large and positive shift and slide
values, indicating that the platinated base is significantly displaced toward the major groove. As
will be discussed in more detail later, this feature is also present in structures of platinated DNA
containing bound proteins and is believed to be a key recognition element for proteins that interact with platinated DNA.
In addition to the structure of the 1,2-intrastrand cross-link, that of the 1,3-intrastrand
cross-link on duplex DNA has also been solved by NMR spectroscopy. 59' 60 This lesion, a likely
major adduct of carboplatin-DNA binding, distorts double-stranded DNA in a different manner
than the 1,2-d(GpG) cross-link (Fig. 1.2.b). In this structure the duplex is bent by ~20' and the
double helix displays local unwinding and widening of the minor groove, similarly to features of
the structure of the 1,2-d(GpG) cross-link. The 1,3-d(GpTpG) adduct differs, however, in that
base pairing of the 5' G*-C, where the asterisk denotes a platination site, is disrupted and the internal thymidine of the adduct is extruded outside the double helix. Although the area of the duplex in the immediate vicinity of the 1,3-d(GpTpG) adduct is more severely distorted than in the
1,2-d(GpG) counterpart, the global effects of the 1,3-cross-link on the DNA duplex are more
subtle than for the 1,2-lesion, with a less dramatic bend angle.
The structure of a DNA molecule containing a site-specific interstrand cisplatin crosslink was solved both by X-ray crystallography 61 (Fig. 1.2.c) and by NMR spectroscopy. 62 Features of this Pt-DNA adduct are structurally unique in many ways compared to those of the intrastrand cross-links. In the ICL, the {Pt(NH 3)2}2+ moiety binds in the minor groove and bends
the helix by 470 in that direction. The double helix is severely unwound by 1100 and displays
local left-handedness, resulting in the two cytosine bases opposite the bound guanosines to be
pointed outward, away from the duplex. As in the intrastrand cross-links, the Pt-N7 bonds are
strained, with the Pt atom being displaced from the guanine ring planes by 0.3 - 0.6 A. Platinum
ICLs on DNA also adopt a unique head-to-tail binding conformation whereby the ligating guanine bases are oriented in opposite directions. 63 Head-to-head binding is observed in all intrastrand cross-links of cisplatin on double-stranded DNA.
Oxaliplatin. Oxaliplatin produces a similar type of DNA adduct spectrum as cisplatin and carboplatin, although oxaliplatin-DNA lesions contain a {Pt(DACH)}
2+
(DACH = trans-R,R-
diaminocyclohexane) rather than a {Pt(NH 3)2}2+ group. 64' 65 DNA duplexes containing sitespecific 1,2- {Pt(DACH) }2+ -d(GpG) adducts have been studied both by X-ray crystallography 66
and by NMR spectroscopy.6 7 The X-ray crystal structure is very similar to that of the analogous
cisplatin-damaged DNA, with the duplex bent toward the major groove and the double helix taking on an A/B-form hybrid structure (Fig. 1.2.d). The NMR solution structure revealed the oxaliplatin-damaged DNA to be mostly B-form, further emphasizing the effect of crystal packing
on the X-ray structure. The solution structure was overall very similar to that of DNA bearing a
cisplatin 1,2-d(GpG) cross-link, but the global bend angle was only 310, compared to 800 for the
cisplatin adduct.
(a)
(b)
(c)
(d)
(e)
(f)
Figure 1.2. X-ray crystal and NMR structures of double stranded DNA containing adducts of
various platinum anticancer agents. (a) Cisplatin 1,2-d(GpG) intrastrand cross-link (1AIO). (b)
Cisplatin 1,3-d(GpTpG) intrastrand cross-link (1DA4). (c) Cisplatin interstrand cross-link
(lA2E). (d) Oxaliplatin 1,2-d(GpG) intrastrand cross-link (1PG9). (e) Satraplatin 1,2-d(GpG)
intrastrand cross-link (lLU5). (f) Pyriplatin monofunctional adduct (3CO3). PDB accession
codes are given in parentheses.
Despite the similarity of the oxaliplatin-1,2-d(GpG) structure to that of the cisplatin lesion, several conformational differences were observed. 68' 69 The cisplatin cross-link preferentially forms hydrogen bonding interactions on the 5' side of the adduct and causes more structural distortion to the base pair step at the 5' end. Conversely, oxaliplatin forms hydrogen bonds
more readily to the 3' side of the intrastrand cross-link. Particularly pronounced is an interaction
between a hydrogen atom of the NH 2 group of the DACH ligand and the 06 oxygen atom of the
3' guanine base. 66 This interaction can form only with the biologically active R,R-isomer of oxaliplatin and not the inactive S,S-isomer. It has been postulated that these conformational differences between oxaliplatin- and cisplatin-DNA adducts may be responsible for differences in pro68 70
tein recognition and cellular processing of the two platinum antitumor compounds. -
Non-classical platinum compounds. In addition to cisplatin/carboplatin and oxaliplatin, the
DNA adducts of several additional cytotoxic platinum compounds have been structurally characterized. The most clinically relevant of these complexes to be investigated was satraplatin, cc,tammine(cyclohexylamine)dichlorodiacetatoplatinum(IV), a platinum(IV) complex that reached
Phase III clinical trials for treatment of hormone-refractory prostate cancer.'
7
The axial acetate
ligands are released as the platinum complex is reduced in the bloodstream, and the resulting
platinum(II) complex binds DNA in a manner analogous to that of cisplatin. Two orientational
isomers form, in which the cyclohexylamine ligand is pointed either toward the 3' or the 5' end
of the platinated DNA strand. These adducts appear in approximately a 2:1 ratio in favor of the
3' -orientational isomer. The structure of the major isomer of this asymmetric bifunctional 1,2d(GpG) adduct was
characterized
crystallographically
on a dodecamer
duplex. cis-
Ammine(cyclohexylamine)platinum(II)-DNA adducts derived from satraplatin cause the same
conformational changes to the double helix as other platinum 1,2-d(GpG) cross-links (Fig.
1.2.e).714
Several multi-platinum-containing complexes have been studied and the structures of the
resulting DNA adducts characterized. A bifunctional cross-link of the bis-platinum compound
[{trans-PtC(NH 3)2} 2 (H2N(CH 2)4NH2)]2+ was studied on self-complementary octamer DNA in
solution by NMR spectroscopy.75 This non-traditional platinum complex showed cytotoxicity in
a series of cisplatin-resistant cell lines, 76 and forms a variety of intra- and inter-strand cross-links
made possible by the extended butanediamine linker between DNA-binding moieties.77 '78 Instead
of forming double-stranded DNA in solution, a unique hairpin structure assembled with two
DNA strands connected via one platinum complex in a head-to-head conformation, forming an
overall dumbbell structure. This unanticipated result underscores the fact that this platinum complex is capable of forming a vastly different spectrum of adducts compared to cisplatin and its
analogs.
[PtCl(en)(ACRAMTU-S)](N0 3)2 (en=ethane-1,2-diamine, ARAMTU=1-[2-(acridin-9-ylamino)ethyl]-1,3-dimethylthiourea) is a dual metalating/intercalating DNA binding drug conjugate that shows cytotoxicity in a range of solid tumor cell lines. 79 -' In 80% of its adducts, this
complex binds to guanine-N7 in the major groove, selectively at d(CpG) sites. 82 This adduct was
characterized on octamer duplex DNA by NMR spectroscopy.83 The modified sequence shows
structural features reminiscent of both B- and A-type DNA. Platinum is bound to the N7 position
of guanine in the major groove, and the intercalating agent is inserted into the d(CpG) base-pair step
on the 5'-face of the platinated nucleobase. The Pt lesion lengthens (rise, 6.62
A) and
unwinds
(twist, 15.40) the duplex at the central base-pair step but does not cause helical bending, which distinguishes PT-ACRAMTU-induced damage from the 1,2-intrastrand cross-link formed by cisplatin. DNA adducts of this complex inhibit binding of the TATA-binding protein to its promoter,
suggesting a mechanism by which Pt-ACRAMTU could inhibit transcription. 2
Another multiple-Pt containing complex [{trans-Pt(NH3)2(NH2(CH 2)(NH 3 )
2 -U-{trans-
Pt(NH 3 )2 -(NH 2 (CH 2 )6 NH 2)2 }], or TriplatinNC, was crystallized on the Dickerson B-DNA dodecamer8 4 and structurally characterized. 85 This active compound is unique in that it does not contain labile leaving groups; instead it binds DNA non-covalently through hydrogen-bonding interactions between the Pt-N ligands and phosphate oxygen atoms in a bidentate N--O---N fashion,
called the "Phosphate Clamp." TriplatinNC selectively binds oxygen atoms in the phosphodiester
backbone, and causes a modest bend in the DNA helix. Whereas TriplatinNC is an active but not
clinical compound, an analog of this complex, BBR3464, contains one chloride ligand on each of
the terminal Pt atoms, and is currently in Phase II clinical trials. It was proposed that BBR3464
may interact transiently with DNA via the Phosphate Clamp prior to binding at the purine N7
atoms.
A cationic platinum(II) complex containing three inert amine ligands and only one labile
leaving group, cis-diammine(pyridine)chloroplatinum(II) (pyriplatin or cDPCP), has antitumor
activity in animal tumor models 23 and in cell lines. 8 6 The X-ray crystal structure of this complex
on duplex DNA is discussed in Chapter 2 and provided the first geometric information about an
antitumor active monofunctional platinum-DNA adduct (Fig. 1.2.fi).2387 This complex inhibits
transcription at a level comparable to that of cisplatin as revealed by in vitro studies. 86 Like oxaliplatin, it is selectively taken up by cells bearing organic cation transporters (OCTs), 86' 88 which
presents an opportunity for delivery to colorectal tumor cells that express OCT membrane proteins in high abundance. 88' 89 The global structure of cDPCP-damage DNA is quite different from
that of DNA containing a platinum intrastrand d(GpG) cross-link. 86'90 The latter platinated duplex is bent by ~40' toward the major groove at the site of the cross-link, yet the monofunctional
platinum-dG lesion causes no significant distortion of the double helix. Like the cisplatin intrastrand cross-link, however, the monofunctional adduct creates a distorted base pair step to the
5' side of the platinum site that may be correlated to antitumor activity.86 Table 1.1 summarizes
features of the various Pt-DNA adducts that have been structurally characterized to date.
Effects of platinum binding on nucleosome structure. In eukaryotic organisms, ~80% of genomic DNA is wrapped in nucleosomes, which consists of 146 base pairs of DNA wrapped in a
left-handed superhelix around a core of eight histone proteins. 9 1,92 It is therefore necessary to
consider this component of the cellular environment when studying the interactions of platinum
compounds with their biological target. Biochemical methods have been used to study the structural effects of cisplatin binding on nucleosome structure and dynamics. 93-95 Hydroxyl radical
and exonuclease footprinting revealed that cisplatin intrastrand cross-links direct nucleosome
positioning to a preferred rotational setting, with the {Pt(NH 3)2}2 moiety pointing inward toward the histone octamer protein core. This preferred position overrides that of strong native
DNA positioning sequences and occurs in nucleosomes prepared from both native, containing a
variety of post-translational modifications, and recombinant histones. Other studies demonstrated
that cisplatin or oxaliplatin adducts inhibit ATP-independent nucleosome mobility in samples of
nucleosome core particles treated with either drug. 96 These data demonstrate that platinum complexes influence not only the structure of the DNA double helix, but also that of nucleosomes.
Chapters 5 and 6 describe new studies exploring the effects of cisplatin binding on nucleosomal
structure and dynamics.
Unwinding
(deg)
20
Pt
location
Major
groove
DNA form
Reference
X-ray
Bend angle
(deg)
35-40
A/B
53, 54
12
NMR
22
20
Major
groove
B
69
Cisplatin
1,2-(GG) intrastrand
(HMGB1-bound)
16
X-ray
45
21
Major
groove
B
120
Cisplatin
1,3-(GTG) intrastrand
13
NMR
20
19
Major
groove
B
59
Cisplatin
(GC/GC) interstrand
10
X-ray
47
70
Minor
groove
Z-like
61
Cisplatin
(GC/GC) interstrand
10
NMR
20
87
Minor
groove
Z-like
62
Oxaliplatin
1,2-(GG) intrastrand
12
X-ray
30
20
Major
groove
A/B
66
Oxaliplatin
1,2-(GG) intrastrand
12
NMR
31
25
Major
groove
B
69
Satraplatin
1,2-(GG) intrastrand
12
X-ray
38
20
Major
groove
A/B
74
Pyriplatin
dG monofunctional
12
X-ray
N/A
8
Major
groove
B
86, 90
TriplatinNC
"Phosphate Clamp"
12
X-ray
16
N/A
DNA
backbone
B
85
DNA length
(bp)
12
Method
Cisplatin
1,2-(GG) intrastrand
Pt adduct
Cisplatin
1,2-(GG) intrastrand
Table 1.1. Structural features of platinum-DNA adducts characterized by NMR spectroscopy and X-ray crystallography.
Protein Binding to Platinum-DNA Adducts
A number of proteins have been identified that bind to Pt-DNA adducts with specificity
over unmodified DNA, including those associated with DNA repair, HMG-domain proteins,
transcription factors, and others.9 7 -99 Within the scope of this thesis, only proteins that play a role
in eukaryotic transcription will be discussed. Transcription factors that bind Pt-DNA include
human upstream binding factor (hUBF), TATA-binding protein (TBP), and Y-box binding protein (YB-1). High-mobility group box protein 1 (HMGB 1), an abundant non-histone chromosomal protein, binds cisplatin-DNA adducts tightly and with selectivity. HMGB1 is implicated to
play a role in the mechanism of action of cisplatin in a variety of ways. Structure specific recognition protein 1 (SSRP1), an HMG-domain containing protein, is a subunit of FACT (facilitates
chromatin transcription), which is a critical chromatin remodeling factor involved in transcription of nucleosomal DNA. SSRP 1 was the one of the first HMG-domain proteins known to bind
platinated DNA. PARP-1, a multi-functional protein with many roles in cells, controls transcriptional levels,100'010 binds selectively to cisplatin-DNA cross-links, and is activated in response to
platinum treatment of cells.
Upstream binding factor. The interaction between HMG-domain proteins and Pt-DNA adducts
has been thoroughly studied. 1o,1o3 One member of this class of proteins, the ribosomal RNA
transcription factor hUBF, binds the cisplatin 1,2-d(GpG) cross-link with a Kd of 60 pM, the
highest known affinity of any protein toward a Pt-DNA lesion.104 In an in vitro transcription assay with RNA polymerase I, treatment of DNA with cisplatin inhibited ribosomal RNA synthesis
051 06
by sequestering hUBF.1 '
TATA-binding protein. The TATA-binding protein is a critical transcription factor for all three
eukaryotic RNA polymerases (pol I, II, and III).107 This protein binds DNA at promoter sites in
the minor groove, bending the double helix toward the major groove and causing a structural
change similar to that of a cisplatin intrastrand cross-link (see Fig. 1.3a).10 8 TBP binding to the
1,2- {Pt(NH 3 )2 }2+-d(GpG) adduct is similar to that of a promoter binding in terms of affinity, with
Kd~ 0.3-10 nM. The kinetics are also similar, with relatively slow on and off rates. TBP binds
the 1,2-d(GpG) cross-link of cisplatin better than the 1,3-d(GpTpG) adduct.109
Y-box binding protein. Another transcription factor that binds cisplatin-modified DNA is YB- 1,
a protein that recognizes an inverted CCAAT sequence termed the Y-box." 0 This protein is
important both for signaling of DNA damage and for cell proliferation. YB-1 binds selectively to
1,2-d(GpG), 1,2-d(ApG), and 1,3-d(GpTpG) cross-links of cisplatin,in and is overexpressed in
the nuclei of cisplatin-resistant cell lines.11 2,113 mRNA for YB-1 is increased approximately 6fold as a response to cisplatin treatment."
4
High-mobility group box protein 1. HMGB1 has been implicated to have a regulatory effect on
many cellular processes involving DNA, including chromatin remodeling, recombination, replication, and transcription.1'151 16 The relationship between HMGB1 levels and cisplatin sensitivity
is reviewed elsewhere.1 2 Interest in the role of HMGB 1, and HMG-domain proteins in general, in
mediating cisplatin cytotoxicity has stimulated much research into the interactions between the
HMG domain and Pt-DNA adducts. HMGB 1 contains two tandem HMG domains, A and B, and
a C-terminal acidic tail. The binding affinity of domain A for the 1,2-{Pt(NH 3)2 }2+-d(GpG) adduct depends on the flanking nucleotide sequence, with Kd values ranging between 1.6 - 517 nM.
The range of binding affinities for domain B is slightly weaker, between 48 - 1300 nM." 7 The
full-length protein binds the cisplatin intrastrand cross-link primarily through the A domain with
a dissociation constant of 120 nM." 8 HMGB1 also recognizes the interstrand cross-link of cisplatin, with approximately 5-fold lower affinity.1 19 The structure of a complex between a 16mer
duplex DNA containing a centralized 1,2-d(GpG) cisplatin intrastrand cross-link and the A domain of HMGB1 was solved by X-ray crystallography (Fig. 1.3b).12 0 The HMG domain binds
the adduct in the widened minor groove to the 3' side of the platinated strand. A phenylalanine
residue intercalates into a hydrophobic notch created by the cisplatin cross-link; binding of the
domain is dramatically reduced when this residue is mutated to alanine. These data provide insight into the recognition of Pt-DNA adducts by all HMG-domain-containing and other proteins.
(a)(b
Figure 1.3. Protein recognition and binding to Pt-DNA adducts. (a) Overlay of X-ray crystal
structures of TBP-bound DNA (1TGH, blue) and DNA containing a cisplatin 1,2-d(GpG) intrastrand cross-link (1AIO, burgundy). (b) X-ray crystal structure of HMGB1 domain A bound to
a cisplatin 1,2-d(GpG) intrastrand cross-link (1CKT). An intercalated phenylalanine residue
plays a key role in substrate recognition. PDB accession codes are given in parentheses.
Structure specific recognition protein 1. SSRP1 was discovered from expression screening of
a human B-cell cDNA library as a protein that binds to cisplatin modified DNA.1 03 This protein,
along with Sptl6, comprise the FACT heterodimer, which alleviates the nucleosomal barrier to
transcription.!" FACT binds cisplatin globally-modified DNA and the 1,2-d(GpG) cross-link
with specificity over undamaged DNA or DNA treated with trans-diamminedichloroplatinum(II).
2
Isolated SSRP1 did not form a high-affinity complex with cisplatin-DNA ad-
ducts, demonstrating the requirement for Sptl6 in recognition of the platinum damage, but the
truncated HMG domain of SSRP1 did recognize the 1,2-d(GpG) cross-link. The affinity of this
critical transcriptional mediator for cisplatin-DNA damage suggests that binding of SSRP1 and
FACT to platinum cross-links may be important to the mechanism of transcription inhibition by
this drug.
Poly(ADP-ribose) polymerase 1. PARP-1 catalyzes the addition of ADP-ribose moieties from
NAD* to the carboxyl groups of protein residues. 123 PARP-1 is implicated in regulation of transcription both by modification of histones to affect chromatin structure and nucleosome stability,124 ,12 5 and by acting on transcriptional enhancers and promoters. 12 6 In addition to being activated by DNA damage agents, 12 6 PARP-1 binds selectively to the cisplatin 1,2-d(GG) intrastrand
12 7
cross-link. 97' 98 In many cases inhibition of PARP-1 sensitizes cell lines to cisplatin treatment.
Inhibition of PARP-1 catalysis also resulted in an increase of protein binding to cisplatin-DNA
adducts.128 Combination treatment of tumors with cisplatin and PARP inhibitors is currently being evaluated in Phase I and II clinical trials. 129
Inhibition of Transcription by Platinum Antitumor Complexes
In L1210 leukemia cells, G2 arrest is required for apoptosis, and loss of DNA replication
viability does not correlate with cell death. 13 0-13 2 These observations are a key indication that
transcription inhibition by cisplatin-DNA adducts is critical to programmed cell death. Prior to
these results, inhibition of DNA replication had been widely considered to be a key to the
mechanism of cisplatin cytotoxicity.
33 -'s
The later data suggested that cells arrested in G2 phase
because they could not synthesize mRNA necessary to pass into mitosis, implicating transcription inhibition as a critical determinant in the pathway of apoptosis triggered by cisplatin. Since
these reports, numerous systems employing both site-specifically and globally platinated DNA
templates, with both recombinant proteins and in living cells, have been designed to study inhibition of transcription by cisplatin and other platinum anticancer agents. Taken together, the data
clearly demonstrate that the ability of a platinum complex to block RNA synthesis correlates directly with its efficacy as an antitumor agent.136
Reconstituted systems. Initial studies of transcription inhibition by platinum antitumor agents
utilized DNA containing site-specific Pt adducts transcribed by purified mammalian RNA polymerase II (pol II) and E. coli RNA polymerase (RNAP). 137-140 Data from these experiments
demonstrated that 1,2-d(GpG) and 1,2-d(ApG) adducts of cisplatin blocked both polymerases
almost completely when placed on the DNA template strand, whereas transcription was only
slightly inhibited when the lesions were placed on the non-template strand. The 1,2{Pt(NH 3)2}2 -d(GpG) cross-link reduced binding affinity of E. coli RNAP and increased the apparent K.. of the enzyme by a factor of 4-5.138 Furthermore, 1,3-d(GpTpG) cross-links of both
cis- and trans-diamminedichloroplatinum(II) strongly blocked elongation by both RNA polymerases.141 Modest inhibition was also observed when the 1,3-cross-link was located on the nontemplate, or coding, strand. Bifunctional Pt cross-links were much more effective at impeding
transcription progression than monofunctional cisplatin adducts. Furthermore, arrested transcrip-
tion elongation complexes were identified as substrates of the RNA transcript cleavage reaction
mediated by TFIIS, indicating that the stalled elongation complex is not released from template
DNA. 14 0 Other studies using globally platinated DNA probes and T7 or SP6 RNA polymerases
showed that transcription was halted primarily at 1,2-d(GpG) or d(ApG) Pt adduct sites, and to a
lesser extent at the cisplatin ICL locations, 142 but no inhibition was observed due to monofinctional adducts of [Pt(dien)Cl]* or cis-[Pt(NH 3)2(H2O)Cl]*.
14 3
Interstrand cross-links of trans-
DDP were similarly effective at blocking these enzymes.144
Use of an immobilized DNA template allows for a high degree of control over transcriptional experiments. Such systems have been utilized in more recent investigations of RNA polymerase inhibition by Pt-DNA adducts to provide additional mechanistic insight. In the first of
these reports, site-specific 1,2-Pt-d(GpG) and 1,3-Pt-d(GpTpG) adducts of both cisplatin and oxaliplatin were incorporated into DNA strands that were subjected to both promoter-dependent
and -independent transcription by T7 RNAP in a reconstituted system. 14 5 All four adducts
strongly block transcription by the enzyme, with the oxaliplatin 1,3-d(GpTpG) adduct providing
the greatest inhibition, followed by cisplatin 1,3-d(GpTpG), cisplatin 1,2-d(GpG), then oxaliplatin 1,2-d(GpG) cross-links in decreasing order. It was also discovered that UTP is incorrectly incorporated into the RNA strand opposite the platinated guanosine residue and that stalled
polymerases can resume transcriptional activity upon removal of the platinum adduct by cyanide
treatment.
Studies in cell extracts and cell culture. Other investigations of platinated DNA templates were
performed either in live cells or using cell extracts. The first such report utilized a plasmid containing a p-galactosidase (p-gal) reporter gene transfected into HeLa, CHO, or human lym-
phoblastoid cell lines.14 6 Transcriptional activity was monitored colorimetrically by addition of
the p-gal substrate ortho-nitrophenol-p-galactoside. Plasmids treated with cisplatin inhibited
transcription 2-3-fold more readily than plasmids treated with trans-DDP. In this system RNA
pol II bypassed cis- and trans-DDP adducts with efficiencies of 0-16% and 60-70%, respectively, and approximately four-fold more trans-DDPrelative to cisplatin was required to block
gene expression by 63%. Transcription of adenovirus major late promoter containing templates
by RNA pol II in cell extracts was inhibited by treatment with cisplatin in a concentrationdependent manner.14 7 Transcription of an undamaged template was also blocked by the addition
of exogenous platinum-damaged DNA, indicating that platinum adducts may inhibit transcription initiation by hijacking essential transcription factors that bind Pt-DNA adducts. In the same
study it was demonstrated that cisplatin adducts can inhibit transcription elongation as well. Sitespecific 1,2-d(GpG) or 1,3-d(GpTpG) intrastrand cross-links of cisplatin were introduced into
DNA and used as transcription templates. Both adducts were efficient blocks of T3 RNA polymerase, and the 1,3-d(GpTpG) cross-link inhibited transcription elongation by RNA pol II by 80%.
Interestingly, pol II efficiently bypassed the 1,2-d(GpG) lesion, although this bypass may be a
sequence-specific result, in light of other data 1 7 ,140 that demonstrate nearly complete inhibition
of pol II by the cisplatin 1,2-d(GpG) cross-link.
Transcription of immobilized DNA templates containing site-specific Pt-DNA adducts in
HeLa nuclear extracts revealed further details of pol II inhibition.14 8 The arrested enzyme remains stably associated with the Pt damage site and is capable of resuming transcription if the
platinum is removed. In HeLa cell culture, stalled pol II was ubiquitylated by ubiquitin ligases at
Lys6, Lys-48, and Lys-63. However, only some portion of the modified enzyme was released
from the DNA and degraded by proteasomes; the rest remained stably bound at the Pt-DNA adduct site.
Data from other studies measuring transcription fidelity in cells correlate well with those
collected from in vitro experiments. Treatment of human fibroblast cells with 50 p.M cisplatin
resulted in a 45% decrease in mRNA levels and increased expression of p53 and p21 .149 Treatment of mouse tumor cells stably transfected with the mouse mammary tumor virus promoter
(MMTV) with cisplatin resulted in highly inhibited expression. MMTV has a well-characterized
chromatin structure, and these experiments determined that, concomitant with reduced RNA levels, chromatin remodeling and transcription factor binding were also inhibited.15 0 These effects
were not observed when the cells were treated with trans-DDP.Most recently, transcription from
site-specifically platinated plasmids was investigated in live cells using a dual luciferase assay."5
These results showed (i) that a single 1,2-d(GpG) or 1,3-d(GpTpG) cross-link of cisplatin was
capable of blocking transcription nearly completely in live cells, and (ii) that transcriptional fidelity recovered over time in DNA-repair-competent cells, but did not recover in cell lines in
which DNA repair was knocked down.
Transcription-coupled repair of Pt-DNA adducts. Transcription-coupled repair (TCR) is a
sub-pathway of nucleotide excision repair (NER) that allows DNA damage sites recognized by
stalled RNA polymerases to be preferentially removed.15 2 TCR deficiency in cells has been positively correlated with cisplatin sensitivity, whereas cells lacking proteins for global NER exhibit
typical levels of resistance to platinum treatment.15 3 1,
4
If transcription inhibition is a critical de-
terminant of cytotoxicity by platinum drugs, then the mechanism by which Pt-DNA adducts
elude transcription-coupled repair must be investigated. However, the TCR pathway in mammal-
ian cells is not well understood, and there is much still to be learned about the mechanism of
TCR and its role in processing Pt-DNA damage.
The connection between transcription inhibition by cisplatin damage and DNA repair was
investigated using immobilized DNA
templates."sis
A site-specific 1,3-{Pt(NH 3 )2}2 -
d(GpTpG) intrastrand cross-link was incorporated into the template DNA to provide an absolute
block to transcription by pol II. The fate of the stalled polymerase and repair of the cisplatin lesion were then examined both in whole cell extracts and in a reconstituted system. In repairproficient extracts, the Pt-DNA adduct was removed by dual excision without release of pol I.155
The elongation complex stalled at the damage site was stable to detergent washes, but could be
removed in an ATP-dependent process. RNA polymerase II containing a dephosphorylated carboxyl-terminal domain was more sensitive to release. In the reconstituted system, the stalled
elongation complex recruited several repair proteins, including TFIIH, XPA, RPA, XPG, and
XPF, in an ATP-dependent manner. 156 In the presence of CSB, the platinum lesion was excised
and the RNA polymerase partially released.
RNA polymerase II is ubiquitylated in cells in response to transcription inhibition by cisplatin or UV-damage, or a-amanitin treatment. 141'15 This effect is not observed in cells deficient
in TCR,i58 and has been demonstrated in both live cells and nuclear extracts. These results are
consistent with ubiquitylation being an important step in the recognition of stalled pol II elongation complexes. Ubiquitylated pol II was partially released from template DNA and degraded by
the proteasome, but the rest remained stably bound at the arrest site. The consequence of pol II
removal by this mechanism has been debated: one possibility is that polymerase removal is required to allow access of repair proteins to the damage site. Another possibility is that degradation of the stalled polymerase triggers an alternative pathway to TCR.
The Mechanism of Transcription Inhibition by Pt-DNA Adducts
The evidence to date has shown that DNA adducts of platinum antitumor compounds inhibit eukaryotic transcription and strongly suggests that this process is directly correlated to its
efficacy as a chemotherapy agent. More recently, effort has been focused on establishing the
mechanism of this process. What is the molecular pathway linking formation of platinum-DNA
adducts to disruption of RNA synthesis? Hypotheses about how cisplatin and its relatives inhibit
transcription can be divided into three categories: (1) hijacking of transcription factors, (2) a
physical block of the enzyme, and (3) inhibition at the stage of chromatin remodeling. These hypotheses are not mutually exclusive; one or more of them may play a role in the cisplatin mechanism of action. Because previous studies suggest that platinum anticancer agents block transcription at both the initiation and elongation stages, it is likely that inhibition occurs by more than
one mechanism. These theories are summarized in Fig. 1.4 and explained in more detail below.
Transcription factor hijacking. According to the transcription factor hijacking hypothesis, PtDNA adducts inhibit RNA synthesis by serving as binding sites for transcription factors such as
the TATA-binding protein that have high affinity for platinated DNA. Interactions with cisplatin
adducts prevent these transcription factors from binding their native promoter sites, thus inhibiting transcription at the initiation stage. The strongest evidence for this theory comes from the
observation that transcription of an undamaged DNA plasmid in human cell extracts can be inhibited in a concentration-dependent manner by introduction of an exogenous cisplatin-modified
DNA substrate. 147 Furthermore, it was demonstrated that microinjection of TBP into living cells
in which transcription levels had been reduced by either cisplatin- or UV-damage resulted in reversal of the inhibition. 159 A similar but less dramatic effect was observed after introduction of
the basal transcription factors TFIIB and TFIIH. Analysis of the X-ray crystal structures of the
TATA-TBP complex160 and double-stranded DNA containing the 1,2-d(GpG) intrastrand crosslink of cisplatin reveals strong similarities between the structures of the double helix in each
model (Fig. 1.3.a).159 The bifunctional platinum adduct creates a bent DNA structure that mimics its protein-bound form. Together these data collectively suggest that transcription factor hijacking by platinum-DNA adducts prevents the assembly of transcription elongation complexes
at promoter sites and inhibits the initiation of RNA synthesis.
Pt
Pt
Transcriptionfactor
hijacking
Roadblock to RNA polymerases
Disruption of chromatin
structure/mobility
Figure 1.4. Possible mechanisms of transcription inhibition by platinum antitumor agents. Platinum-modified DNA can recruit transcription factors to the damage site, preventing these proteins
from binding promoter sites and blocking formation of elongation complexes. The Pt-DNA adduct can also serve as a physical block to RNA polymerases when the lesion is located on the
transcribed DNA strand. Finally, Pt-DNA adducts can disrupt nucleosomal structure and/or mobility and block transcription by prohibiting access to DNA by transcriptional proteins.
Roadblock of RNA polymerases. Data from many in vitro transcription systems indicate that
14 14 5 156
platinum-DNA adducts inhibit RNA polymerases at the site of the cisplatin cross-link, 0' '
suggesting that the DNA adducts serve as a physical impediment to transcription elongation by
the enzyme. Recently an X-ray crystal structure of RNA pol II stalled at a cisplatin 1,2-d(GpG)
intrastrand cross-link revealed how the DNA damage may inhibit the enzyme.161 In this structure
the Pt adduct is located downstream of the pol II active site, in the +2/+3 positions along the
template strand; ribonucleotide addition occurs at the +1 position of the template. Attempts to
place the cross-link inside the active site of the elongation complex resulting in backtracking of
the polymerase so that the damage site was again located downstream of the reaction site. This
result indicates that the adduct is not stably accommodated in the +1/+2 or -1/+1 positions. From
these data and from analysis of the X-ray crystal structure, the authors proposed that the cisplatin
cross-link inhibits pol II due to a translocation barrier whereby the bases in the platinated
dinucleotide cannot rotate properly to allow entry into to the enzyme active site, stalling the protein upstream of the platinum damage site. Biochemical experiments demonstrated that the elongation complex misincorporates AMP across from the 3' guanosine of the Pt lesion. The kinetics
and manner of this process are consistent with established nontemplated AMP incorporation
known as the "A-rule," 162 which provides more evidence that the Pt cross-link never enters the
pol II active site. Stalling is independent of the G-A mismatch, indicating that the translocation
barrier arising from the Pt cross-link, not the mismatch, is primarily responsible of pol II inhibition. If the intrastrand cross-link is introduced and transiently stabilized upstream of the proposed
translocation barrier, then lesion bypass and revived transcription occurs.
Disruption of chromatin dynamics. Another possible manner by which cisplatin-DNA adducts
may interfere with transcription could occur at the chromatin level. Proper nucleosomal positioning and mobility are critical to the fidelity of transcription.163,64 Initial transcription factor binding occurs at DNA promoter sites that are characteristically nucleosome-free, which allows the
proteins to recognize and bind the naked DNA sequence. As the RNA polymerase elongation
complex subsequently transcribes along the template DNA, upstream nucleosomes are continu-
ally shifted and unwrapped by chromatin remodeling complexes such as FACT.121 , 164 Data on the
effects of cisplatin intrastrand cross-links on nucleosome positioning and mobility suggest that
platinum damage can inhibit transcription at both the initiation and elongation stages by altering
the nucleosomal organization of promoter sites and reducing nucleosome mobility, respectively.
Nucleosome positioning is determined primarily by the intrinsic DNA sequence, 91"165 but
site-specific 1,2-d(GpG) or 1,3-d(GpTpG) intrastrand cross-links of cisplatin enforce a characteristic rotational positioning of the DNA strand around the nucleosome, such that the Pt adduct
faces inward towards the histone core. 93 -95 This effect overrides that of native DNA positioning
sequences and could disrupt native nucleosomal organization and potentially disturb protein recognition of binding sites by placing a nucleosome at the promoter position if it occurs in vivo.
Other studies suggest that platinum damage does not significantly effect nucleosome positioning, but rather reinforces the native conformation. In these experiments nucleosome core
particles were treated with cisplatin or oxaliplatin, and both electrophoretic mobility shift assays
and X-ray crystallographic studies of crystals of NCP treated with the drugs indicated no structural changes. 9 6,166 The difference between the two sets of experiments is that the former results
describe formation of nucleosomes from site-specifically platinated DNA, where the position and
structure of the adducts are known, whereas the latter involve global treatment of nucleosomes
with cisplatin or oxaliplatin, which will provide a heterogeneous panoply of adducts. These results suggest that platinum binds nucleosomal DNA in positions where adducts are most readily
accommodated by the nucleosome structure, thus reinforcing the native positioning preference
instead of modifying it.
Platinum intrastrand cross-links disrupt nucleosomal dynamics as well as structure. Nucleosomes treated with cisplatin or oxaliplatin exhibit significantly decreased heat-induced mo-
bility. Inhibited nucleosomal sliding could also limit transcription by preventing access of the
RNA polymerase to the DNA template. A similar mechanism has been proposed for a series of
pyrrole-imidazole polyamides that bind nucleosomal DNA, inhibit nucleosomal mobility, and
block transcription from a nucleosomal template, but not from naked DNA.167-'
These polyam-
ides reduce nucleosome sliding through a proposed blockage of DNA rotation around the histone
octamer. Data from this system suggest that inhibition of nucleosome mobility is sufficient to
reduce transcriptional activity. Given that platinum intrastrand cross-links cause a similar reduction in DNA sliding on the nucleosome, cisplatin may also block transcription through this twist
diffusion mechanism. Chapters 5 and 6 describe new studies of the effects of cisplatin-DNA
damage on nucleosome structure and dynamics.
From Transcription Inhibition to Apoptosis
Many transcriptional inhibitors have been tested as antitumor agents, including compounds that bind directly to RNA polymerases 171 ,172 and agents that block transcription by inhibiting phosphorylation of pol II by CDK9.173 , 174 Inhibition of transcription induces a cellular response leading to the activation of p53, a tumor suppressor protein, through the ATR
kinase.149 1 75 Induction of p53 connects blocked RNA polymerase with cell cycle checkpoints,
DNA repair, and apoptosis. After a certain time point, if the transcription block persists the cells
will undergo apoptosis in either a p53-dependent or -independent manner. The mechanism of this
process is not clearly understood, but several pathways have been proposed, as reviewed in detail
elsewhere 17 5"17 6 and briefly discussed here. In general, the half-lives of mRNAs encoding for proapoptotic proteins such as Bax or Bid are longer than for anti-apoptotic proteins such as Bcl-2 or
Mcl-i; thus inhibition of transcription over an extended period of time may adjust the ratios of
pro- and anti-apoptotic proteins to conditions favoring cell death. 77',"' Transcription inhibition
may also lead to apoptosis through either p53-dependent or -independent pathways. The role of
p53 in this process is highly controversial; the p53-dependent pathway may involve translocation
of the protein to mitochondria so that it can bind anti-apoptotic Bcl-2 family proteins and induce
cell death.' 7 9 However, other evidence suggests that transcription blockage can induce apoptosis
in cells lacking p53.iso- 82 A third possibility is that stalled transcription elongation complexes
can block the replication machinery if the polymerase is not removed from the template DNA
prior to cell entry into S phase. Finally, export of certain proteins from the cell nucleus requires
constant synthesis and export of mRNAs.'
83
It is therefore possible that apoptosis may result
from inability to shuttle key proteins from the nucleus to the cytoplasm.
The proposed pathways from inhibition of transcription to apoptosis are mentioned in the
context of platinum antitumor drug mechanism because a common characteristic of tumor cells
is an impaired ability to undergo apoptosis.184 Resistance of certain cancers to cisplatin has been
thoroughly investigated and many mechanisms are in play.185 However, one aspect often ignored
by the community is that a functional signaling pathway to apoptosis is critical to the efficacy of
platinum anticancer compounds.
Future Directions
Much progress has been made in elucidating the mechanism of action of cisplatin, one of
the most successful anticancer therapeutics to date. With this information in hand, researchers
can begin to take a rational approach to designing new platinum complexes that specifically target the pathways involved. Traditionally the focus of platinum drug design has been to prepare
compounds that form intrastrand DNA cross-links like cisplatin. In the future, research should
focus not on only on forming certain types of DNA adducts, but on targeting and manipulating a
cellular pathway triggered by the DNA damage. In recent years, the entire field of cancer research has moved away from general cytotoxic agents and focused more on targeted therapies,
tissue. 186-189
where delivered agents accumulate preferentially in the tumor and spare healthy
Many strategies have been devised to synthesize tumor-specific platinum complexes. 190-194 By
combining transcription-inhibiting design with tumor-specific accumulation, researchers should
be able to significantly improve the ability of platinum complexes to treat cancer.
References
(1)
Loehrer, P. J.; Einhorn, L. H. Ann. Intern. Med. 1984, 100, 704-713.
(2)
Keys, H. M.; Bundy, B. N.; Stehman, F. B.; Muderspach, L. I.; Chafe, W. E.; Suggs III,
C. L.; Walker, J. L.; Gersell, D. New Eng. J. Med. 1999, 340, 1154-1161.
(3)
Morris, M.; Eifel, P. J.; Lu, J.; Grigsby, P. W.; Levenback, C.; Stevens, R. E.; Rotman,
M.; Gershenson, D. M.; Mutch, D. G. New Eng. J. Med. 1999, 340, 1137-1143.
(4)
Bosl, G. J.; Bajorin, D. F.; Sheinfeld, J.; Motzer, R. J.; Chaganti, R. S. K. In Cancer:
Principles & practice of oncology; 6th ed.; DeVita Jr., V. T., Hellman, S., Rosenberg, S.
A., Eds.; Lippincott Williams & Wilkins: Philadelphia, 2001, p 1491-1518.
(5)
Bosl, G. J.; Motzer, R. J. New Eng. J. Med. 1997, 337, 242-253.
(6)
Kartalou, M.; Essigmann, J. M. MutationRes. 2001, 478, 23-43.
(7)
Canetta, R.; Bragman, K.; Smaldone, L.; Rozencweig, M. Cancer Treat. Rev. 1988, 15,
17-32.
(8)
Raymond, E.; Faivre, S.; Chaney, S.; Woynarowski, J.; Cvitkovic, E. Mol. Cancer Ther.
2002, 1, 227-235.
(9)
Rosenberg, B.; Van Camp, L.; Krigas, T. Nature 1965, 205, 698-699.
(10)
Rosenberg, B.; Van Camp, L.; Trosko, J. E.; Mansour, V. H. Nature 1969, 222, 385-386.
(11)
Wang, D.; Lippard, S. J. Nat. Rev. DrugDiscovery 2005, 4, 307-320.
(12)
Jung, Y.; Lippard, S. J. Chem. Rev. 2007, 107, 1387-1407.
(13)
Gale, G. R.; Morris, C. R.; Atkins, L. M.; Smith, A. B. CancerRes. 1973, 33, 813-818.
(14)
Binks, S. P.; Dobrota, M. Biochem. Pharmacol. 1990, 40, 1329-1336.
(15)
Gately, D. P.; Howell, S. B. Br. J. Cancer 1993, 67, 1171-1176.
(16)
Ishida, S.; Lee, J.; Thiele, D. J.; Herskowitz, I. Proc. Natl. Acad. Sci. U.S.A. 2002, 99,
14298-14302.
(17)
Holzer, A. K.; Samimi, G.; Katano, K.; Naerdemann, W.; Lin, X.; Safaei, R.; Howell, S.
B. Mol. Pharmacol.2004, 66, 817-823.
(18)
Howe-Grant, M. E.; Lippard, S. J. In Metal Ions Biol. Syst.; Sigel, H., Ed.; Marcel
Dekker: New York, 1980; Vol. 11, p 63-125.
(19)
Cohen, G. L.; Ledner, J. A.; Bauer, W. R.; Ushay, H. M.; Caravana, C.; Lippard, S. J. J.
Am. Chem. Soc. 1980, 102, 2487-2488.
(20)
Cleare, M. J.; Hoeschele, J. D. Bioinorg. Chem. 1973, 2, 187-210.
(21)
Farrell, N. In Metal Ions Biol. Syst.; Sigel, A., Sigel, H., Eds.; Marcel Dekker: New York,
2004; Vol. 42, p 251-296.
(22)
Kelland, L. R.; Abel, G.; McKeage, M. J.; Jones, M.; Goddard, P. M.; Valenti, M.;
Murrer, B. A.; Harrap, K. R. CancerRes. 1993, 53, 2581-2586.
(23)
Hollis, L. S.; Amundsen, A. R.; Stem, E. W. J. Med. Chem. 1989, 32, 128-136.
(24)
Perez, J. M.; Fuertes, M. A.; Alonso, C.; Navarro-Ranninger, C. Crit. Rev. Oncol.
Hematol. 2000, 35, 109-120.
(25)
Aris, S. M.; Farrell, N. P. Eur. J. Inorg. Chem. 2009, 2009, 1293-1302.
(26)
Caradonna, J. P.; Lippard, S. J.; Gait, M. J.; Singh, M. J. Am. Chem. Soc. 1982, 104,
5793-5795.
(27)
Fichtinger-Schepman, A. M. J.; Van der Veer, J. L.; Den Hartog, J. H. J.; Lohman, P. H.
M.; Reedijk, J. Biochemistry 1985, 24, 707-713.
(28)
Eastman, A. Biochemistry 1986, 25, 3912-3915.
(29)
Terheggen, P. M. A. B.; Floot, B. G. J.; Scherer, E.; Begg, A. C.; Fichtinger-Schepman,
A. M. J.; den Engelse, L. CancerRes. 1987, 47, 6719-6725.
(30)
Mantri, Y.; Lippard, S. J.; Baik, M.-H. J. Am. Chem. Soc. 2007, 129, 5023-5030.
(31)
Blommaert, F. A.; van Dijk-Knijnenburg, H. C. M.; Dijt, F. J.; den Engelse, L.; Baan, R.
A.; Berends, F.; Fichtinger-Schepman, A. M. J. Biochemistry 1995, 34, 8474-8480.
(32)
Lepre, C. A.; Strothkamp, K. G.; Lippard, S. J. Biochemistry 1987, 26, 5651-5657.
(33)
Eastman, A.; Jennerwein, M. M.; Nagel, D. L. Chem. Biol. Interact. 1988, 67, 71-80.
(34)
Pascoe, J. M.; Roberts, J. J. Biochem. Pharmacol.1974, 23, 1345-1357.
(35)
DeNeve, W.; Valeriote, F.; Tapazoglou, E.; Everett, C.; Khatana, A.; Corbett, T. Invest.
New Drugs 1990, 8, 17-24.
(36)
Scovell, W. M.; Kroos, L. R. Biochem. Biophys. Res. Comm. 1982, 104, 1597-1603.
(37)
Cohen, G. L.; Bauer, W. R.; Barton, J. K.; Lippard, S. J. Science 1979, 203, 1014-1016.
(38)
Macquet, J.-P.; Butour, J.-L. Biochimie 1978, 60, 901-914.
(39)
Nunomura, K.; Maeda, Y.; Ohtsubo, E. J. Gen. Appl. Microbiol. 1991, 37, 207-214.
(40)
Kagemoto, A.; Takagi, H.; Naruse, K.; Baba, Y. Thermochim. Acta 1991, 190, 191-201.
(41)
Maeda, Y.; Nunomura, K.; Ohtsubo, E. J. Mol. Biol. 1990, 215, 321-329.
(42)
Naser, L. J.; Pinto, A. L.; Lippard, S. J.; Essigmann, J. M. Biochemistry 1988, 27, 43574367.
(43)
Poklar, N.; Pilch, D. S.; Lippard, S. J.; Redding, E. A.; Dunham, S. U.; Breslauer, K. J.
Proc.Natl. Acad. Sci. U.S.A. 1996, 93, 7606-7611.
(44)
Pilch, D. S.; Dunham, S. U.; Jamieson, E. R.; Lippard, S. J.; Breslauer, K. J. J. Mol. Biol.
2000, 296, 803-812.
(45)
Rice, J. A.; Crothers, D. M.; Pinto, A. L.; Lippard, S. J. Proc. Natl. Acad. Sci. US.A.
1988, 85, 4158-4161.
(46)
Bellon, S. F.; Lippard, S. J. Biophys. Chem. 1990, 35, 179-188.
(47)
Bellon, S. F.; Coleman, J. H.; Lippard, S. J. Biochemistry 1991, 30, 8026-8035.
(48)
Sip, M.; Schwartz, A.; Vovelle, F.; Ptak, M.; Leng, M. Biochemistry 1992, 31, 25082513.
(49)
Malinge, J.-M.; Perez, C.; Leng, M. Nucleic Acids Res. 1994, 22, 3834-3839.
(50)
Sherman, S. E.; Gibson, D.; Wang, A. H.-J.; Lippard, S. J. Science 1985, 230, 412-417.
(51)
Admiraal, G.; Van der Veer, J. L.; De Graaff, R. A. G.; Den Hartog, J. H. J.; Reedijk, J.
J. Am. Chem. Soc. 1987, 109, 592-594.
(52)
Sherman, S. E.; Gibson, D.; Wang, A. H.-J.; Lippard, S. J. J. Am. Chem. Soc. 1988, 110,
7368-7381.
(53)
Takahara, P. M.; Rosenzweig, A. C.; Frederick, C. A.; Lippard, S. J. Nature 1995, 377,
649-652.
(54)
Takahara, P. M.; Frederick, C. A.; Lippard, S. J. J. Am. Chem. Soc. 1996, 118, 1230912321.
(55)
Yang, D.; van Boom, S. S. G. E.; Reedijk, J.; van Boom, J. H.; Wang, A. H.-J.
Biochemistry 1995, 34, 12912-12920.
(56)
Gelasco, A.; Lippard, S. J. Biochemistry 1998, 37, 9230-9239.
(57)
Marzilli, L. G.; Saad, J. S.; Kuklenyik, Z.; Keating, K. A.; Xu, Y. J. Am. Chem. Soc.
2001, 123, 2764-2770.
(58)
Sullivan, S. T.; Saad, J. S.; Fanizzi, F. P.; Marzilli, L. G. J. Am. Chem. Soc. 2002, 124,
1558-1559.
(59)
Van Garderen, C. J.; Van Houte, L. P. A. Eur. J. Biochem. 1994, 225, 1169-1179.
(60)
Teuben, J.-M.; Bauer, C.; Wang, A. H.-J.; Reedijk, J. Biochemistry 1999, 38, 1230512312.
(61)
Coste, F.; Malinge, J.-M.; Serre, L.; Shepard, W.; Roth, M.; Leng, M.; Zelwer, C. Nucleic
Acids Res. 1999, 27, 1837-1846.
(62)
Huang, H.; Zhu, L.; Reid, B. R.; Drobny, G. P.; Hopkins, P. B. Science 1995, 270, 18421845.
(63)
Sullivan, S. T.; Ciccarese, A.; Fanizzi, F. P.; Marzilli, L. G. J Am. Chem. Soc. 2001, 123,
9345-9355.
(64)
Jennerwein, M. M.; Eastman, A.; Khokhar, A. Chem. Biol. Interact. 1989, 70, 39-49.
(65)
Woynarowski, J. M.; Chapman, W. G.; Napier, C.; Herzig, M. C. S.; Juniewicz, P. Mol.
Pharmacol.1998, 54, 770-777.
(66)
Spingler, B.; Whittington, D. A.; Lippard, S. J. Inorg. Chem. 2001, 40, 5596-5602.
(67)
Wu, Y.; Pradhan, P.; Havener, J.; Boysen, G.; Swenberg, J. A.; Campbell, S. L.; Chaney,
S. G. J. Mol. Biol. 2004, 341, 1251-1269.
(68)
Sharma, S.; Gong, P.; Temple, B.; Bhattacharyya, D.; Dokholyan, N. V.; Chaney, S. G. J.
Mol. Biol. 2007, 373, 1123-1140.
(69)
Wu, Y.; Bhattacharyya, D.; King, C. L.; Baskerville-Abraham, I.; Huh, S.-H.; Boysen,
G.; Swenberg, J. A.; Temple, B.; Campbell, S. L.; Chaney, S. G. Biochemistry 2007, 46,
6477-6487.
(70)
Chaney, S. G.; Campbell, S. L.; Bassett, E.; Wu, Y. Crit.Rev. Oncol. Hematol. 2005, 53,
3-11.
(71)
Kelland, L. R. Exp. Opin. Invest. Drugs 2000, 9, 1373-1382.
(72)
Sternberg, C. N.; Whelan, P.; Hetherington, J.; Paluchowska, B.; Slee, P. H. T. J.;
Vekemans, K.; Van Erps, P.; Theodore, C.; Koriakine, 0.; Oliver, T.; Lebwohl, D.;
Debois, M.; Zurlo, A.; Collette, L. Oncology 2005, 68, 2-9.
(73)
Hartwig, J. F.; Lippard, S. J. J. Am. Chem. Soc. 1992, 114, 5646-5654.
(74)
Silverman, A. P.; Bu, W.; Cohen, S. M.; Lippard, S. J. J. Biol. Chem. 2002, 277, 4974349749.
(75)
Yang, D.; Boom, S. v.; Reedijk, J.; Boom, J. v.; Farrell, N.; Wang, A. H.-J. Nat. Struct.
Biol. 1995, 2, 577-578.
(76)
Farrell, N.; Qu, Y.; Feng, L.; Van Houten, B. Biochemistry 1990, 29, 9522-9531.
(77)
Johnson, A.; Qu, Y.; Van Houten, B.; Farrell, N. Nucleic Acids Res. 1992, 20, 1697-1703.
(78)
Zou, Y.; Van Houten, B.; Farrell, N. Biochemistry 1994, 33, 5404-5410.
(79)
Martins, E. T.; Baruah, H.; Kramarczyk, J.; Saluta, G.; Day, C. S.; Kucera, G. L.;
Bierbach, U. J. Med. Chem. 2001, 44, 4492-4496.
(80)
Hess, S. M.; Anderson, J. G.; Bierbach, U. Bioorg. Med. Chem. Lett. 2004, 15, 443-446.
(81)
Hess, S. M.; Mounce, A. M.; Sequeira, R. C.; Augustus, T. M.; Ackley, M. C.; Bierbach,
U. Cancer Chemotherapy andPharmacology2005, 56, 337-343.
(82)
Budiman, M. E.; Bierbach, U.; Alexander, R. W. Biochemistry 2005, 44, 11262-11268.
(83)
Baruah, H.; Wright, M. W.; Bierbach, U. Biochemistry 2005, 44, 6059-6070.
(84)
Drew, H. R.; Wing, R. M.; Takano, T.; Broka, C.; Tanaka, S.; Itakura, K.; Dickerson, R.
E. Proc.Natl. Acad. Sci. US.A. 1981, 78, 2179-2183.
(85)
Komeda, S.; Moulaei, T.; Woods, K.; Chikuma, M.; Farrell, N.; Williams, L. J. Am.
Chem. Soc. 2006, 128, 16092-16103.
(86)
Lovejoy, K. S.; Todd, R. C.; Zhang, S.; McCormick, M. S.; D'Aquino, J. A.; Reardon, J.
T.; Sancar, A.; Giacomini, K. M.; Lippard, S. J. Proc. Natl. Acad Sci. U.S.A. 2008, 105,
8902-8907.
(87)
Hollis, L. S.; Sundquist, W. I.; Burstyn, J. N.; Heiger-Bemays, W. J.; Bellon, S. F.;
Ahmed, K. J.; Amundsen, A. R.; Stern, E. W.; Lippard, S. J. CancerRes. 1991, 51, 18661875.
(88)
Zhang, S.; Lovejoy, K. S.; Shima, J. E.; Lagpacan, L. L.; Shu, Y.; Lapuk, A.; Chen, Y.;
Komori, T.; Gray, J. W.; Chen, X.; Lippard, S. J.; Giacomini, K. M. Cancer Res. 2006,
66, 8847-8857.
(89)
Hayer-Zillgen, M.; Brass, M.; B6nisch, H. Br. J. Pharmacol.2002, 136, 829-836.
(90)
Todd, R. C.; Lippard, S. J. In Platinum and Other Heavy Metal Compounds: Molecular
Mechanisms and ClinicalApplications; Bonetti, A., Howell, S. B., Leone, R., Muggia, F.,
Eds.; Humana Press: Totowa, 2009, p 67-72.
(91)
Segal, E.; Fondufe-Mittendorf, Y.; Chen, L.; Thistrim, A.; Field, Y.; Moore, I. K.;
Wang, J.-P. Z.; Widom, J. Nature 2006, 442, 772-778.
(92)
Kornberg, R. D.; Lorch, Y. Nat. Struct. Mol. Biol. 2007, 14, 986-988.
(93)
Danford, A. J.; Wang, D.; Wang,
U.S.A. 2005, 102, 12311-12316.
(94)
Ober, M.; Lippard, S. J. J. Am. Chem. Soc. 2007, 129, 6278-6286.
(95)
Ober, M.; Lippard, S. J. J. Am. Chem. Soc. 2008, 130, 2851-2861.
Q.; Tullius,
T. D.; Lippard, S. J. Proc. Natl. Acad. Sci.
(96)
Wu, B.; Davey, C. A. Chem. Biol. 2008, 15, 1023-1028.
(97)
Zhang, C. X.; Chang, P. V.; Lippard, S. J. J. Am. Chem. Soc. 2004, 126, 6536-6537.
(98)
Guggenheim, E. R.; Xu, D.; Zhang, C. X.; Chang, P. V.; Lippard, S. J. Chem. Biochem.
2009, 10, 141-157.
(99)
Zhu, G.; Lippard, S. J. Biochemistry 2009, 48, 4916-4925.
(100)
Kraus, W. L.; Lis, J. T. Cell 2003, 113, 677-683.
(101)
Aguilar-Quesada, R.; Munoz-Gamez, J. A.; Martin-Oliva, D.; Peralta-Leal, A.; QuilesPerez, R.; Rodriguez-Vargas, J. M.; Ruiz de Almodovar, M.; Conde, C.; Ruiz-Extremera,
A.; Oliver, F. J. Curr. Med. Chem. 2007, 14, 1179-1187.
(102)
Scovell, W. M.; Muirhead, N.; Kroos, L. R. Biochem. Biophys. Res. Commun. 1987, 142,
826-835.
(103)
Bruhn, S. L.; Pil, P. M.; Essigmann, J. M.; Housman, D. E.; Lippard, S. J. Proc. Natl.
Acad. Sci. U.S.A. 1992, 89, 2307-2311.
(104)
Treiber, D. K.; Zhai, X.; Jantzen, H.-M.; Essigmann, J. M. Proc. Natl. Acad. Sci. U.S.A.
1994, 91, 5672-5676.
(105)
Jordan, P.; Carmo-Fonseca, M. Nucleic Acids Res. 1998, 26, 2831-2836.
(106) Zhai, X.; Beckmann, H.; Jantzen, H.-M.; Essigmann, J. M. Biochemistry 1998, 37,
16307-16315.
(107)
Cormack, B. P.; Struhl, K. 1992, 69, 685-696.
(108)
Coin, F.; Frit, P.; Viollet, B.; Salles, B.; Egly, J.-M. Mol. Cell BIol. 1998, 18, 3907-3914.
(109)
Jung, Y.; Mikata, Y.; Lippard, S. J. J. Biol. Chem. 2001, 276, 43589-43596.
(110)
Didier, D. K.; Schiffenbauer, J.; Woulfe, S. L.; Zacheis, M.; Schwartz, B. D. Proc. Natl.
Acad. Sci. US.A. 1988, 85, 7322-7326.
(111)
Ise, T.; Nagatani, G.; Imamura, T.; Kato, K.; Takano, H.; Nomoto, M.; Izumi, H.;
Ohmori, H.; Okamoto, T.; Ohga, T.; Uchiumi, T.; Kuwano, M.; Kohno, K. Cancer Res.
1999, 59, 342-346.
(112)
Ohga, T.; Koike, K.; Ono, M.; Makino, Y.; Itagaki, Y.; Tanimoto, M.; Kuwano, M.;
Kohno, K. CancerRes. 1996, 56, 4224-4228.
(113)
Yahata, H.; Kobayashi, H.; Kamura, T.; Amada, S.; Hirakawa, T.; Kohno, K.; Kuwano,
M.; Nakano, H. J. CancerRes. Clin. Oncol. 2002, 128, 621-626.
(114)
Uramoto, H.; Izumi, H.; Ise, T.; Tada, M.; Uchiumi, T.; Kuwano, M.; Yasumoto, K.;
Funa, K.; Kohno, K. J.Biol. Chem. 2002, 277, 31694-31702.
(115)
Baxevanis, A. D.; Landsman, D. Nucl. Acids Res. 1995, 23, 1604-1613.
(116)
Thomas, J. 0.; Travers, A. A. Trends Biochem. Sci. 2001, 26, 167-174.
(117)
Dunham, S. U.; Lippard, S. J. Biochemistry 1997, 36, 11428-11436.
(118)
Jung, Y.; Lippard, S. J. Biochemistry 2003, 42, 2664-2671.
(119) Kasparkova, J.; Delalande, 0.; Stros, M.; Elizondo-Riojas, M.-A.; Vojtiskova, M.;
Kozelka, J.; Brabec, V. Biochemistry 2003, 42, 1234-1244.
(120)
Ohndorf, U.-M.; Rould, M. A.; He, Q.; Pabo, C. 0.; Lippard, S. J. Nature 1999, 399, 708712.
(121)
Reinberg, D.; Sims III, R. J. J. Biol. Chem. 2006, 281, 23297-23301.
(122)
Yarnell, A. T.; Oh, S.; Reinberg, D.; Lippard, S. J. J.Biol. Chem. 2001, 276, 2573625741.
(123)
Schreiber, V.; Dantzer, F.; Ame, J.-C.; de Murcia, G. Nat. Rev. Mol. Cell Biol. 2006, 7,
517-528.
(124) Huletsky, A.; de Murcia, G.; Muller, S.; Hengartner, M.; Menard, L.; Lamarre, D.;
Poirier, G. G. J.Biol. Chem. 1989, 264, 8878-8886.
(125)
Poirier, G. G.; de Murcia, G.; Jongstra-Bilen, J.; Niedergang, C.; Mandel, P. Proc. NatL.
Acad. Sci. U.S.A. 1982, 79, 3423-3427.
(126)
D'Amours, D.; Desnoyers, S.; D'Silva, I.; Poirier, G. G. Biochem. J. 1999, 342, 249-268.
(127)
Miknyoczki, S. J., Jones-Bolin, S., Pritchard, S., Hunter, K., Zhao, H., Wan, W., Ator,
M., Bihovsky, R., Hudkins, R., Chatterjee, S., Klein-Szanto, A., Dionne, C., Ruggeri, B.
Mol Cancer Ther 2003, 2, 371-382.
(128)
Guggenheim, E. R.; Ondrus, A. E.; Movassaghi, M.; Lippard, S. J. Bioorg. Med. Chem.
2008, 16, 10121-10128.
(129)
Helleday, T.; Petermann, E.; Lundin, C.; Hodgson, B.; Sharma, R. A. Nat. Rev. Cancer
2008, 8, 193-204.
(130)
Sorenson, C. M.; Eastman, A. CancerRes. 1988, 48, 4484-4488.
(131)
Sorenson, C. M.; Eastman, A. CancerRes. 1988, 48, 6703-6707.
(132)
Sorenson, C. M.; Barry, M. A.; Eastman, A. J. Natl. Cancer Inst. 1990, 82, 749-755.
(133)
Harder, H. C.; Smith, R. G.; Leroy, A. F. CancerRes. 1976, 36, 3821-3829.
(134)
Johnson, N. P.; Hoeschele, J. D.; Kuemmerle, N. B.; Masker, W. E.; Rahn, R. 0. Chem.
Biol. Interact. 1978, 23, 267-271.
(135)
Pinto, A. L.; Lippard, S. J. Proc.NatL. Acad. Sci. U.S.A. 1985, 82, 4616-4619.
(136)
Sandman, K. E.; Marla, S. S.; Zlokarnik, G.; Lippard, S. J. Chem. Biol. 1999, 6, 541-55 1.
(137)
Corda, Y.; Job, C.; Anin, M.-F.; Leng, M.; Job, D. Biochemistry 1991, 30, 222-230.
(138)
Corda, Y.; Anin, M.-F.; Leng, M.; Job, D. Biochemistry 1992, 31, 1904-1908.
(139)
Corda, Y.; Job, C.; Anin, M.-F.; Leng, M.; Job, D. Biochemistry 1993, 32, 8582-8588.
(140)
Tomaletti, S.; Patrick, S. M.; Turchi, J. J.; Hanawalt, P. C. J. Bio. Chem. 2003, 278,
35791-35797.
(141)
Larsen, E.; Kwon, K.; Coin, F.; Egly, J.-M.; Klungland, A. DNA Repair 2004, 3, 14571468.
(142)
Lemaire, M.-A.; Schwartz, A.; Rahmouni, A. R.; Leng, M. Proc. Nati. Acad. Sci. US.A.
1991, 88, 1982-1985.
(143)
Brabec, V.; Boudnf', V.; Balcarova, Z. Biochemistry 1994, 33, 1316-1322.
(144)
Brabec, V.; Leng, M. Proc. NatL. Acad Sci. US.A. 1993, 90, 5345-5349.
(145)
Jung, Y.; Lippard, S. J. J. Biol. Chem. 2003, 278, 52084-52092.
(146)
Mello, J. A.; Lippard, S. J.; Essigmann, J. M. Biochemistry 1995, 34, 14783-14791.
(147)
Cullinane, C.; Mazur, S. J.; Essigmann, J. M.; Phillips, D. R.; Bohr, V. A. Biochemistry
1999, 38, 6204-6212.
(148)
Jung, Y.; Lippard, S. J. J. Biol. Chem. 2006, 281, 1361-1370.
(149)
Ljungman, M.; Zhang, F.; Chen, F.; Rainbow, A. J.; McKay, B. C. Oncogene 1999, 18,
583-592.
(150)
Mymryk, J. S.; Zaniewski, E.; Archer, T. K. Proc.NatL. Acad Sci. U.S.A. 1995, 92, 20762080.
(151)
Ang, W. H.; Myint, M. N. Z.; Lippard, S. J. J. Am. Chem. Soc. 2010, submitted.
(152)
Svejstrup, J. Q. Nat. Rev. Mol. Cell Biol. 2002, 3, 21-29.
(153)
Furuta, T.; Ueda, T.; Aune, G.; Sarasin, A.; Kraemer, K. H.; Pommier, Y. Cancer Res.
2002, 62, 4899-4902.
(154)
Bulmer, J. T.; Zacal, N. J.; Rainbow, A. J. Cancer Chemother. Pharmacol. 2005, 56,
189-198.
(155)
Tremeau-Bravard, A.; Riedl, T.; Egly, J.-M.; Dahmus, M. E. J. Biol. Chem. 2004, 279,
7751-7759.
(156)
Laine, J.-P.; Egly, J.-M. EMBO J. 2006, 25, 387-397.
(157)
Lee, K.-B.; Wang, D.; Lippard, S. J.; Sharp, P. A. Proc.Nati. Acad. Sci. U.S.A. 2002, 99,
4239-4244.
(158)
Bregman, D. B.; Halaban, R.; van Gool, A. J.; Henning, K. A.; Friedberg, E. C.; Warren,
S. L. Proc.NatL. Acad. Sci. U.S.A. 1996, 93, 11586-11590.
(159)
Vichi, P.; Coin, F.; Renaud, J.-P.; Vermeulen, W.; Hoeijmakers, J. H. J.; Moras, D.; Egly,
J.-M. EMBO J. 1997, 16, 7444-7456.
(160) Juo, Z. S.; Chiu, T. K.; Leiberman, P. M.; Baikalov, I.; Berk, A. J.; Dickerson, R. E. J.
Mol. Bio. 1996, 261, 239-254.
(161)
Damsma, G. E.; Alt, A.; Brueckner, F.; Carell, T.; Cramer, P. Nat. Struct. Mol. Biol.
2007, 14, 1127-1133.
(162)
Strauss, B. S. BioEssays 1991, 13, 79-84.
(163)
Luger, K. Chromosome Res. 2006, 14, 5-16.
(164) Workman, J. L. Genes Dev. 2006, 20, 2009-2017.
(165)
Kaplan, N.; Moore, I. K.; Fondufe-Mittendorf, Y.; Gossett, A. J.; Tillo, D.; Field, Y.;
LeProust, E. M.; Hughes, T. R.; Lieb, J. D.; Widom, J.; Segal, E. Nature 2009, 458, 362366.
(166)
Wu, B.; Dr6ge, P.; Davey, C. A. Nat. Chem. Biol. 2008, 4,110-112.
(167)
Gottesfeld, J. M.; Melander, C.; Suto, R. K.; Raviol, H.; Luger, K.; Dervan, P. B. J. Mol.
Biol. 2001, 309, 615-629.
(168)
Gottesfeld, J. M.; Belitsky, J. M.; Melander, C.; Dervan, P. B.; Luger, K. J. Mol. Biol.
2002, 321, 249-263.
(169)
Suto, R. K.; Edayathumangalam, R. S.; White, C. L.; Melander, C.; Gottesfeld, J. M.;
Dervan, P. B.; Luger, K. J.Mol. Biol. 2003, 326, 371-380.
(170)
Edayathumangalam, R. S.; Weyermann, P.; Gottesfeld, J. M.; Dervan, P. B.; Luger, K.
Proc. Nati. Acad. Sci. U.S.A. 2004, 101, 6864-6869.
(171)
Sobell, H. M. Proc.Nati. Acad. Sci. U.S.A. 1985, 82, 5328-533 1.
--
l-
(172)
Bushnell, D. A.; Cramer, P.; Kornberg, R. D. Proc. Natl. Acad. Sci. U.S.A. 2002, 99,
1218-1222.
(173)
Blagosklonny, M. V. Cell Cycle 2004, 3, 1537-1542.
(174)
Radhakrishnan, S. K.; Gartel, A. L. CancerRes. 2006, 66, 3264-3270.
(175)
Derheimer, F. A.; Chang, C.-W.; Ljungman, M. Eur. J. Cancer 2005, 41, 2569-2576.
(176)
Gartel, A. L. Biochim. Biophys. Acta 2008, 1786, 83-86.
(177)
Lam, L. T.; Pickeral, 0. K.; Peng, A. C.; Rosenwald, A.; Hurt, E. M.; Giltnane, J. M.;
Averett, L. M.; Zhao, H.; Davis, R. E.; Sathyamoorthy, M.; Wahl, L. M.; Harris, E. D.;
Mikovits, J. A.; Monks, A. P.; Hollingshead, M. G.; Sausville, E. A.; Staudt, L. M.
Genome Biol. 2001, 2, 1-11.
(178)
Yang, E.; van Nimwegen, E.; Zavolan, M.; Rajewsky, N.; Schroeder, M.; Magnasco, M.;
Darnell Jr., J. E. Genome Res. 2003, 13, 1863-1872.
(179)
Marchenko, N. D.; Zaika, A.; Moll, U. M. J.Biol. Chem. 2000, 275, 16202-16212.
(180)
Shapiro, G. I.; Koestner, D. A.; Matranga, C. B.; Rollins, B. J. Clin. CancerRes. 1999, 5,
2925-2938.
(181)
Alonso, M.; Tamasdan, C.; Miller, D. C.; Newcomb, E. W. Mol. Cancer Ther. 2003, 2,
139-150.
(182)
Demidenko, Z. N.; Blagosklonny, M. V. CancerRes. 2004, 64, 3653-3660.
(183)
O'Hagan, H. M.; Ljungman, M. Exp. Cell Res. 2004, 297, 548-559.
(184) Hanahan, D.; Weinberg, R. A. Cell 2000, 100, 57-70.
(185)
Kartalou, M.; Essigmann, J. M. Mutation Research 2001, 478, 23-43.
(186) Hait, W. N. CancerRes. 2009, 69, 1263-1267.
(187)
Gordon, A. N.; Tonda, M.; Sun, S.; Rackoff, W. Gynecol. Oncol. 2004, 95, 1-8.
(188)
Gradishar, W. J.; Tjulandin, S.; Davidson, N.; Shaw, H.; Desai, N.; Bhar, P.; Hawkins,
M.; O'Shaughnessy, J. J.Clin. Oncol. 2005, 23, 7794-7803.
(189)
Collins, I.; Workman, P. Nat. Chem. Biol. 2006, 2, 689-700.
(190)
Rice, J. R.; Gerberich, J. L.; Nowotnik, D. P.; Howell, S. B. Clin. Cancer Res. 2006, 12,
2248-2254.
(191)
Kelland, L. Nat. Rev. Cancer 2007, 7, 573-584.
(192) Dhar, S.; Gu, F. X.; Langer, R.; Farokhzad, 0. C.; Lippard, S. J. Proc. NatL. Acad. Sci.
U.S.A. 2008, 105, 17356-17361.
(193)
Dhar, S.; Liu, Z.; Thomale, J.; Dai, H.; Lippard, S. J. J. Am. Chem. Soc. 2008, 130,
11467-11476.
(194) Rieter, W. J.; Pott, K. M.; Taylor, K. M. L.; Lin, W. J. Am. Chem. Soc. 2008, 130, 1158411585.
Chapter 2. X-Ray Crystal Structure of a Monofunctional Platinum-DNA Adduct, cis{Pt(NH 3)2(pyridine)} 2+Bound to Deoxyguanosine in a Dodecamer Duplex
Research in this chapter has been published in Proc. Natl. Acad. Sci. US.A. 2008, 105, 89028907, and Platinum and Other Heavy Metal Compounds: Molecular Mechanisms and Clinical
Applications. 2009, Totowa: Humana Press; 67-72
Introduction
The propensity of cis-diamminedichloroplatinum(II), (cis-[Pt(NH 3)2 Cl 2] or cisplatin) to
form bifunctional intrastrand cross-links on DNA has been linked to its efficacy as an anticancer
drug.' These platinum-DNA adducts interfere with cellular processes such as transcription and
replication and, if left unrepaired, lead to apoptosis. 2 From early animal studies of platinum
compound efficacy, the following structure-activity relationships were developed: 1) the squareplanar platinum(II) complex should contain two labile leaving groups in a cis configuration, 2)
the leaving groups should be -3.4
A apart
(the distance between adjacent bases in B-form DNA),
3) the compound should be neutral to facilitate passive diffusion across the cell membrane, and
4) the inert ligands should be strongly-bonded ammine-type moieties. 3 For years these guidelines
directed the field of platinum antitumor research, leading to the synthesis of thousands of close
structural analogues of cisplatin.
Over time many "rule-breakers" were discovered that violate these structure-activity
relationships, yet show activity in various tumor models. Polynuclear platinum 4 and transplatinum 5 complexes are now being examined in clinical trials, despite their inability to form 1,2intrastrand adducts. The non-classical platinum compound cis-diammine(pyridine)chloroplatinum(II) (pyriplatin or cDPCP, see Fig. 2.1) is a cationic, monofunctional platinum(II)
complex, the activity of which was established almost 20 years ago,6 but has not yet been
pursued in the clinic. This compound, which contains only one leaving group, blocks DNA
replication at single dG sites in replication mapping experiments.! Immunochemical and other
experiments demonstrate that cDPCP forms a fundamentally different adduct than that of
cisplatin, i.e. it does not lose a pyridine or ammine ligand to form a bifunctional cross-link.7'8
Further investigation into the properties of this complex revealed that 1) pyriplatin is selectively
taken up into cells by the organic cation transporter OCT 1, a transporter expressed in colorectal
tumors and a primary avenue for uptake of oxaliplatin,9 (2) adducts of cDPCP, like those of
cisplatin,10'
serve as an effective block of transcription by RNA polymerase II, and (3) cDPCP-
DNA lesions are inefficiently repaired by mammalian NER machinery relative to cisplatin-DNA
cross-links.' 2 Combining these results, it is evident that cDPCP may be an exciting new drug
candidate for treatment of colon cancer.
H3N
HPt CI""PtI
H3N/
\C,
O
H2
NO
C
\0CI
'N
H2
cisplatin
oxaliplatin
OC
0
..- N
NH31
PtNH
/\NH3
cDPCP
Figure 2.1. Chemical structures of cisplatin, oxaliplatin, and pyriplatin.
One major question still to be answered about this complex is why it displays
cytotoxicity, when the vast majority of monofunctional platinum(II) complexes, including the
trans-diammine(pyridine)chloroplatinum(II), are inactive. NMR13 and X-ray crystal' 4 structures
of cisplatin-modified DNA reveal significant bending and unwinding of the double helix,
distortions which are proposed to be critical determinants of cisplatin cytotoxicity in vivo.
Because of the monofunctional nature of the cis-{Pt(NH 3)2(pyridine)}2 -dG lesion, it is likely
that a different type of distortion would account for the unique activity of pyriplatin in tumor
cells, but the structure of such an adduct has never been explored crystallographically. To
investigate this issue, a DNA dodecamer duplex with a site-specific adduct of cDPCP bound to a
central deoxyguanosine residue was synthesized and X-ray diffraction studies initiated.'12,
The
sequence of the DNA is similar to that used in previous studies of platinum-DNA duplexes,14'16'17
but modified to allow only one platinum binding site. The 2.17
A resolution structure reveals that
pyriplatin-DNA adducts do not bend double-strand DNA, but some similarities with cisplatin
intrastrand cross-links were observed. Combined with in vitro transcription results already
reported, these findings provide insight into the molecular mechanism of action of a uniquely
active platinum anticancer agent.
Experimental
Materials. Potassium tetrachloroplatinate(II) as obtained as a gift from Engelhard Corporation
(now BASF, Iselin, NJ). Phosphoramidites and other reagents for DNA synthesis were purchased
from Glen Research. Crystallization reagents were obtained from Hampton Research and Sigma.
Calf intestinal phosphatase was purchased from New England Biolabs, and nuclease P1 was
purchased from MP Biomedical. Oligonucleotide synthesis was performed with an Applied
Biosystems 392 DNA/RNA
automated synthesizer. High-pressure liquid chromatography
(HPLC) was carried out on a Waters 600E system controller with a Waters 486 detector or an
Agilent 1200 series instrument. Atomic absorption and UV-Vis spectroscopy were performed
using a Perkin-Elmer AAnalyst 300 system and a HP 8453 UV-visible spectrophotometer,
respectively. Electrospray ionization mass spectrometry (ESI-MS) was performed on an Agilent
1100 series MSD trap.
Synthesis of cis-diammine(pyridine)chloroplatinum(II)
(1). Synthesis of pyriplatin was
accomplished in three steps as described previously 6'"8 (see Scheme 2.1). K2 PtCl4 (1.5 g, 3.6
mmol) was dissolved in 5 mL ddH20 in a 50 mL round bottom flask, and then heated to 60 'C.
KI (2.96 g, 17.8 mmol, 5 equiv dissolved in 3 mL ddH 20) was slowly added to the solution and
stirred at 60'C for 10 min. 1 mL of 30% NH40H (7.7 mmol, 2 equiv) was added to the solution,
The precipitate was collected by vacuum
and a yellow precipitate immediately formed.
filtration, washed with H20, EtOH, and Et2 O, and dried. The total yield of cis-[Pt(NH 3)212] was
1.60 g (3.3 mmol, 92% yield). AgNO 3 (1.12 g, 6.6 mmol) was dissolved in 5 mL ddH 20 in a 50
mL round bottom flask. Pt(NH 3)212 (1.60 g, 3.3 mmol) was added to the solution as a solid, and
the resulting suspension was stirred for 10 min at room temperature. The solid (AgI) was filtered
off, and the solution was heated to 60 'C. A solution of KCl (0.53 g, 7.1 mmol in 2 mL ddH2O)
was slowly added to the flask, resulting in the formation of a yellow precipitate. After stirring
for 10 min, the flask was allowed to cool to ambient temperature, then cooled to 4 'C over 4 h to
The product was collected by vacuum filtration, washed with H2 0,
complete precipitation.
EtOH, and Et 2O, and dried under vacuum. The total yield of cisplatin was 0.86 g (2.9 mmol,
88% yield).The product was characterized by AA spectroscopy by dissolving 5.70 mg cisplatin
in 500 mL water in a volumetric flask, then diluting 5 mL of the solution into 500 mL water by
volumetric pipette. Expected [Pt]: 74.5 mg/L, found [Pt]: 73.0 ± 0.9 mg/L.
Cl 1)excess KI
C1
K2
Pt
CI
60C, 10
I 1)2eqAgNO 3
H3N
in
C_ 2) 2 eq NH40H
\ /
Pt
H3N/
RT, 10 min
-
C1
H3N
Pt
-
2) 2 eq KCI
60OC, 10 min
1eq pyridine inH2
\ /
H3N
3
C
1
[D
H3N_
\
600C, 2 h/RT, 72 h
H3N
/
C1
Pt
CI
Scheme 2.1. Synthesis of cis-diammine(pyridine)chloroplatinum(II) from K2 PtCl4 .
Cisplatin (500.2 mg, 1.67 mmol) was dissolved in 85 mL ddH 20 in a 250 mL round
bottom flask. A 135 pL portion of pyridine (1.67 mmol, 1 equiv) was added, and the suspension
was heated to 60 0 C with stirring for 2 h, then at room temperature for 70 h in the dark. The
suspension was filtered, concentrated to -10 mL, then filtered again to remove unreacted
cisplatin. The filtered solution was dried completely, and then recrystallized three times from
0.1 N HCl, 0.5 N HCl, and MeOH, respectively. A total of 62.7 mg (0.17 mmol) of 1 was
collected (10% yield).
The final product was characterized by ESI-MS and 195 Pt NMR
spectroscopy. ESI-MS [M*]; calc: 343.0 Da, observed: 343.2 Da. 195Pt NMR; found: -2289 ppm
(referenced to H2 PtCl6 at 0 ppm), observed: -2309 ppm (referenced to K2 PtCl 4 at -1628 ppm).
Deoxyoligonucleotide
synthesis
and
purification.
The
deoxyoligonucleotides
5'-
d(CCTCTGCTCTCC)-3' (2) and 5'-d(CCTCTCGTCTCC)-3' (3), along with their respective
complementary strands (4 and 5), were prepared on a 10.0 gmol scale with dimethoxytrityl
(DMT) groups on, using standard phosphoramidite solid support methodologies,19 and purified
by reversed-phase HPLC (see method RCT.001P, Appendix C). After removal of DMT groups
with 80% acetic acid, the purified oligonucleotide was precipitated with isopropanol and desalted
using Waters Sep-Pak C18 cartridges.
Synthesis of G6-platinated oligonucleotide. A 68 mM aqueous solution of 1 was combined
with 1.98 equiv of silver nitrate in the dark for 9 h to activate the platinum complex. After
removal of silver chloride precipitate by centrifugation, a portion of the supernatant containing
1.2 equiv of platinum was allowed to react with a 0.25 mM (2.0 ptmol) aqueous solution of 2 in
the dark at 37 'C overnight. The platinated oligonucleotide (6) was purified by preparative ionexchange HPLC on a Dionex DNAPac PA- 100 column (9 x 250 mm) by method RCT.002S (see
Fig. 2.2, top, and Appendix C). A total of 1.17 pmol product was collected (58%), which was
characterized by ESI-MS (method RCT.010M in Appendix C, see Fig. 2.3) and enzymatic
digestion with nuclease P1 and calf-intestinal phosphatase as previously described. 2"1
Nucleoside ratios were quantitated by HPLC (Fig. 2.4) using method RCT.008A in Appendix
C. ESI-MS; calculated: 3815.1 Da, observed: 3813.9 Da. The Pt/DNA ratio was determined by
AA and UV-Vis spectroscopy to 0.91 ± 0.07.
1400
1200
9
1000
*800
600
*
2
400
200
0L
0
5
10
is
20
25
30
35
40
25
30
35
40
Time (min)
1600
1400
1200
1000
800
600
400
200
0
s
10
15
20
Time (min)
Figure 2.2. HPLC chromatograms of the purification of single- (peak 1, top) and double- (peak
3, bottom) stranded platinated dodecamer DNA.
if$#"
U261M.4"
2.
2W
4
w
1O
O
o
W
sw
lw
sa2c 4 -u
-5
.5
m
lw
me.
me
s
NiW
low
i
-4
"i
"16
'
0
Id W
Figure 2.3. ESI-MS spectra of DNA strands 6 (top) and 4 (bottom). The charged species z = -7,
z = -6, and z = -5 are shown. ESI-MS; calculated: 3815.1 Da and 3785.1 Da, respectively,
observed: 3813.9 Da and 3784.5 Da, respectively.
Synthesis of G7-platinated oligonucleotide. A 6.7 mM aqueous solution of 1 was combined
with 1.98 equiv of silver nitrate in the dark for 9 h to activate the platinum complex. After
removal of silver chloride precipitate by centrifugation, a portion of the supernatant containing
1.2 equiv of platinum was allowed to react with a 0.25 mM (323 mnol) aqueous solution of 3 in
the dark at 37 *C overnight. The platinated oligonucleotide (7) was purified by preparative ionexchange HPLC using the same method. A total of 170 nmol product was collected (53%). The
Pt/DNA ratio was determined by AA and UV-Vis spectroscopy to be 1.16 ± 0.08.
Preparation of Pt-DNA duplexes. Duplex 8, with the Pt adduct at G6, was formed by
combining equimolar portions 4 and 6 (1.0 pmol each) in 1.7 mL total volume of an aqueous
solution containing 50 mM MgCl 2 , 200 mM LiCl, and 100 mM HEPES pH 7.0. In a similar
manner, duplex 9, containing the platinum moiety at position G7, was prepared by combining
150 nmol each of strands 5 and 7. The products were annealed by heating to 80 'C for 2 min then
slowly cooling to 4 'C over 4 h. Double-stranded DNAs were purified by ion-exchange HPLC
by method RCT.003S (see Fig. 2.2, bottom). The final yield of platinated duplexes 8 and 9 were
0.67 pmol and 114 nmol, respectively,
with Pt/DNA ratios of 1.14 ± 0.04 and 1.04
0.17,
respectively.
dG T dG-Pt
dC
d.
05
000
000
500
1000
000
50
0
25 0 3
3500
4000
4500
0
5500
008006
0 043
0 001
13
000
22
0
0 00
$00
1000
6000
150
1600
000
2500
3000
3600
4000
4600
5000
550
MrnAes
Figure 2.4. HPLC traces of 2, 4, and 6 after enzymatic digestion with nuclease P1 and calf
intestinal phosphatase. Peaks were identified as dC (7.3 min), dG (15.4 min), T (17.2 min), dGPt (18.8 min), and dA (26.7 min). 4: Expected peak ratios (G/A/C): 7/4/1. Found: 6.9/4.0/1.0. 2:
Expected peak ratios (C/T/G): 7/4/1. Found: 7.3/4.3/1.0. 6: The dG peak disappears, and the PtdG peak grows in. Expected peak ratios (C/T): 7/4. Found: 6.8/4.0.
Crystallization and X-ray data collection. The hanging drop vapor diffusion method was used
for crystallization, 20'2 ' with initial conditions obtained from a nucleic acid matrix screen Natrix
(Hampton Research). 2 Clusters of small crystals of duplex 8 initially grew at 4 'C from 4 pL
droplets containing 0.2 mM DNA and 0.5X precipitant solution, equilibrated against 1.0 mL of
IX precipitant solution containing 80 mM Mg(OAc) 2 , 50 mM sodium cacodylate pH 6.5, and
30% w/v polyethylene glycol (PEG) 4000. Diffraction-quality single crystals of duplex 9 with
approximate dimensions of 0.2 x 0.2 x 0.1 mm were subsequently grown from precipitant
solutions containing 120 mM Mg(OAc) 2 , 50 mM Na cacodylate pH 6.5, 1 mM spermine, and
28% w/v PEG 4000. Crystal formation occurred in approximately 2-4 weeks. All solutions were
sterile filtered through a 0.22 ptm membrane immediately prior to use.
Crystals of 9 were transferred to a cyroprotectant solution containing 120 mM
Mg(OAc) 2, 50 mM Na cacodylate pH 6.5, 1 mM spermine, 30% w/v PEG 4000, and 15% v/v
glycerol, then mounted on loops and flash frozen directly in liquid nitrogen. Data sets were
collected at 100 K on beamline 24-ID-C at the Advanced Photon Source at Argonne National
Laboratory. An X-ray fluorescence scan was performed to measure the platinum absorption
spectrum, and the wavelength corresponding to the peak absorption energy (1.0719 A) was
selected for single-wavelength anomalous dispersion (SAD) studies. Data were collected on an
ADSC Q315 detector (360 frames, AF = 10, exposure time = 2 s), and processed in HKL2000.
A representative diffraction image from the anomalous data set is shown in Fig. 2.5. Higher
resolution data were later collected on beamline 9-1 of the Stanford Synchrotron Radiation
Laboratory (180 frames, AF = 1', exposure time = 5 s, MAR 325 detector) and processed with
HKL2000.
Collection statistics for both data sets are summarized in Table 2.1.
Structure determination and refinement. SAD phases were calculated using the program
SHARP,2 4 and an initial model containing only the nucleic acid was manually built into the
resulting solvent-flattened electron density map (shown in Fig. 2.5) with the program Coot.2 5
After cycles of rigid-body refinement in Refmac5 26 and manual rebuilding of the model in Coot,
the platinum atom and ligands were included. This model was then subjected to restrained TLS
refinement" against the high-resolution data to 2.17 A, using restraints for the phosphodiester
backbone, sugars, and nucleobases from the Refmac library. Initial TLS parameters were
obtained from the TLS Motion Determination server.2 8 2 9 Sixteen water molecules were added to
locations with appropriate hydrogen bonding distances (<3.5 A) to the DNA and electron density
greater than 1.5c on the 2Fo-F, map. A simulated-annealing composite omit map was calculated
using CNS. 3 0 Final refinement statistics are given in Table 2.1. Geometric parameters were
calculated using the program 3DNA.3 1 Final coordinates for the refined model were deposited
into the Protein Data Bank with accession code 3CO3.
Figure 2.5. (left) X-ray diffraction image of crystals of 9, used for phasing by SAD methods.
Reflections extend out to -2.7 A. (right) Solvent-flattened electron density map of 9, contoured
at la (blue) The anomalous difference map, contoured at 5G, is depicted in red, indicating the
location of the platinum atom.
Table 2.1. Data collection and refinement statistics
Data collection statistics*
SAD
Data set
APS 24-ID-Ct
Beamline
1.072
Wavelength (A)
C222 1
Space group
Unit cell dimensions (A)
45.8
a
66.4
b
c
56.6
High res.
SSRL 9-2
0.984
C222 1
46.4
66.0
56.1
50-2.72
Resolution range (A)
14263
Obs. reflections
2264
Unique reflections
94.2 (73.6)
Completeness
23.2 (6.7)
I/a
7.3 (17.6)
Rmerge§
Refinement statistics
RII
50-2.17
32977
4765
96.0 (76.0)
18.1 (2.7)
10.7 (55.6)
Rfree1
25.4
RMSD bond lengths (A)
0.006
22.5
RMSD bond angles (0)
1.435
Average B-factors (A2)
Platinum ligand
Solvent
42.4
44.4
42.9
*Values in parentheses are for the highest resolution shell.
tADSC Q315 detector, 360 frames, A1 = 10, exposure time = 2 s
IM4AR 325 detector , 180 frames, A1 = 10, exposure time = 5 s
Rmerge - |II( I.
R = YIFo| - |Fe|||Y|Fo|.
R,,e = R obtained for a test set of reflections (5% of diffraction data).
Results
Sample preparation and crystallization. Oligonucleotide platination proceeded cleanly
because only one purine base was available for cDPCP binding. A small amount of multiply
platinated DNA was observed in the HPLC chromatograms, but these products were readily
separated during the purification process. The secondary binding location is presumably the N3
atom of cytosine, which has been shown previously to serve as a platinum coordination site.
Pt/DNA ratios and ESI-MS data confirmed that each DNA strand contains a single cis-
{Pt(NH 3)2(pyridine)} 2+ moiety, and nuclease digestion results indicated that the central
guanosine is the modified base because the free guanosine peak, observed in the chromatograms
of the unplatinated strands, disappeared completely in those of the platinated strands.
Sample 8 was the first compound prepared for crystallization studies, but diffractionquality crystals were never obtained for this molecule. However, the appearance of crystalline
clusters indicated that the conditions, previously used to grow crystals of duplex DNA containing
an oxaliplatin-DNA adduct,16 may be favorable for crystal growth. Duplex 9, containing the cis{Pt(NH 3)2(pyridine)} 2+-dG adduct on the G7 position instead of G6, which presumably had
sufficiently different packing interactions from those of 8, formed nice single crystals.
Unit cell composition and crystal packing. The duplex crystallized in the orthorhombic space
group C222 1, with one molecule in the asymmetric unit and a solvent content of 56%. It should
be noted that the structure could also be solved in the space group P2 1 with two molecules in the
asymmetric unit. However, the decrease in Rmerge was only marginal (9.1% in P2 1 compared to
10.7% in C222 1), and the R and Rfree for refinement were approximately equal, so the higher
symmetry space group was chosen.
Two predominant packing interactions organize the platinated DNA molecules within the
unit cell (see Fig. 2.6). End-to-end packing, which is commonly encountered in B-DNA
structures, is facilitated by hydrogen bonding of deoxyribose moieties of C1 and C12, and
similarly of G13 with G24, in neighboring duplexes, creating a pseudo-continuous double helix
throughout the crystal. Groove-to-groove packing also occurs between molecules, aided in part
by hydrogen-bonding interactions between an ammine ligand on platinum of one duplex and the
phosphate backbone on G16 of an adjacent molecule. Sixteen water molecules were located,
with the most ordered ones residing in the major groove between two adjacent duplexes.
c11
(a)
G14
(b)
C12
3.2
C18
-(G13
A%
-
3.
A
'
31A17
G7*
G23
A
G16
Figure 2.6. End-to-end (a) and groove-to-groove (b) binding interactions between DNA
molecules that contribute to crystal packing in the unit cell. Asterisk indicates platinum binding
site.
Platinated DNA duplex. The Pt-DNA duplex maintains a linear, B-form conformation despite
coordination of platinum to the central dG residue (see Fig. 2.7), and all Watson-Crick hydrogen
bond base pairs throughout the dodecamer are conserved. The double helix is unwound by
approximately 8' in the vicinity of the platination site, in agreement with previous NMR
spectroscopic results of a DNA duplex modified with the 4-Me-pyridine analog of cDPCP.
No
other distortion of the global nucleic acid structure was observed.
The aromatic ligand of the cis-{Pt(NH 3)2(py)}2+-dG adduct is directed toward the 5' end
of the platinated strand, also in accord with previous NMR data on a DNA duplex with a parasubstituted cDPCP analog bound. 3 This orientation facilitates hydrogen bond formation between
the NH 3 ligand trans to pyridine and 06 of the guanosine residue (N-0 distance, 2.8
A).
Interestingly, this hydrogen bond also occurs in Pt-DNA adducts formed by oxaliplatin, (RR)diaminocyclohexaneoxalatoplatinum(II), but presumably not in adducts formed by the inactive
S,S-(DACH) stereoisomer.16, 34 Hydrogen-bonding interactions in cisplatin- and oxaliplatinDNA adducts have been thoroughly studied using NMR spectroscopy and molecular dynamics
simulations, 34 35
, suggesting that they may be involved in differential recognition of these DNA
damage sites by nuclear proteins. The precise role of these interactions in cellular processing of
Pt-DNA adducts has not yet been elucidated; however, current data indicate that these hydrogen
bonds may be important in the mechanism of action of platinum antitumor compounds.3 4 3 s
(a)
(
(b)
ce
C18
G19
(d)
G7*
Figure 2.7. The structures of (a) pyriplatin- and (b) cisplatin-damaged DNA duplexes. Close-up
views of the platinum binding sites for cDPCP and cDDP are shown in (c) and (d), respectively.
Asterisks indicate platinum binding sites. (a) and (c), PDB accession code 3CO3; (b) and (d),
lAIO.
Compared to cisplatin, pyriplatin binding only moderately distorts the structure of double
helical DNA (Fig. 2. 7.a and b). Characteristics of the cisplatin 1,2-intrastrand cross-link include
a large roll angle between the bound guanines (Fig. 2. 7.d), global bending of ~40' towards the
major groove, and local unwinding of the duplex by ~25." These distortions are hypothesized
to inhibit transcription and, if the DNA damage persists, trigger cellular apoptosis.2 The
monofunctional cDPCP adduct does not affect the roll or global bend angle of the DNA duplex,
and it unwinds the helix by only 8'. The Pt atom of cDPCP lies within the guanine plane, as
shown in Fig. 2.7.a. DNA geometric parameters for the cDPCP- and cisplatin-DNA adducts are
compared in Table 2.2.
C6
G19
G19
C6
(AG7
,fG7-
Figure 2.8. Stereoscopic views of the cDPCP-dG adduct on duplex DNA. Bottom: 2F-Fe
electron density map contoured at la.
Table 2.2. Nucleic acid geometric parameters for Pt-DNA adducts (asterisks indicates
platination site).
Base pair"/Base pair step parameters
Base pair
Sx
Sy
Sz
K
Torsion angles'
a>
a
cDPCP-DNA
Base
a
p
y
e
C
X
-109
-102
-107
cDPCP-DNA: Pt strand (listed 5'-3')
5 T-A
-0.2
-0.1
0.1
-0.2
-18.8
-1.2
5T
-28
161
41
147
-160
6 C-G
0.2
-0.1
0.1
-4.6
-14.2
-1.0
6C
-49
168
37
123
-165
-102
7 G*-C
0.1
-0.1
-0.1
-3.9
-13.1
-0.8
7 G*
-40
167
33
143
-169
-150
-89
8 T-A
-0.1
0.1
0.1
-2.1
-15.1
-1.0
8T
-35
167
37
147
-164
-111
-108
complementary strand (listed 3'-5')
cisplatin-DNA
20 A
-10
155
17
144
-168
-99
-99
5 T-A
0.6
-0.2
0.1
7.1
-3.4
3.5
19 G
-54
157
17
161
-167
-149
-89
6 G*-C
-0.1
-0.3
0.3
15.4
-18.4
-5.5
18 C
-66
172
42
135
-103
173
-99
7 G*-C
-1.0
-0.6
0.2
-2.0
-14.5
-4.6
17 A
-42
160
42
149
-173
-92
-107
8 T-A
0.5
0.2
0.2
-2.4
-22.7
6.9
bp step
Dx
Dy
Dz
r
5 TC/GA
0.4
0.3
3.5
2.2
1.3
39.4
6 CG*/CG
1.2
0.8
3.4
5.8
1.7
33.3
7 G*T/AC
-0.9
-0.5
3.3
-2.6
2.9
31.0
cisplatin-DNA: Pt strand (listed 5'-3')
QQ
cDPCP-DNA
cisplatin-DNA
5T
-67
169
56
77
-173
-80
-154
6G*
-66
7G*
-77
175
52
91
-144
-56
-147
164
73
82
-168
-82
8T
-74
-169
159
62
92
-163
-71
-154
complementary strand (listed 3'-5') '
20 A
151
-169
173
86
-160
-71
-168
19 C
-71
-177
53
83
-162
-68
-157
5 TG*/CA
-1.4
-1.4
3.2
-0.6
10.9
25.6
18 C
-75
176
56
86
-161
-84
-156
6 G*G*/CC
1.5
-1.9
3.4
0.1
26.9
24.8
17 A
-82
149
62
83
-166
-57
-159
7 G*T/AC
-0.2
-1.0
3.4
1.9
2.4
43.1
aBase
pair parameters defined as follows: Sx, shear (A); Sy, stretch (A); Sz, stagger (A);
K,
buckle
(0); c>, propeller twist (0); a, opening (0).
Base pair step parameters defined as follows: Dx, shift (A); Dy, slide (A); Dz, rise (A); r, tilt (0);
g, roll (0); Q, twist (0).
cTorsion angles (0) are defined as Phos-a-05'-pi-C5'-y-C4'-6-C3'-e-03'--Phos.
X is the glycosyl
torsion angle.
Although pyriplatin and cisplatin modify nucleic acids in a mono- and bi-functional
manner, respectively, the resulting adducts share a common feature. Pt-DNA damage causes
distortion of the base pair step on the 5' side of the lesion, regardless of the nature of binding.
This bp step (depicted in Fig. 2.8) is marked by large shift and slide values; i.e., the base pair
containing the platinum adduct is translocated out towards the major groove. The respective shift
and slide values are 1.2 A and 0.8 A for the cDPCP-dG adduct, and 1.5 and 1.9 A for the
cisplatin 1,2-intrastrand cross-link.
13
Discussion
Several features of platinum-DNA cross-links are shared by the monofunctional adduct
of cDPCP. First, a hydrogen bonding interaction between the NH 3 ligand cis to guanine and the
06 atom of the nucleobase mimics a similar stereospecific interaction in oxaliplatin-DNA
adducts that has been thoroughly studied using NMR spectroscopic studies and molecular
dynamics simulations.3 4 3 s This hydrogen bond formation, which occurs more readily in
oxaliplatin-DNA adducts than in cisplatin-DNA adducts, may be responsible for conformational
differences that ultimately result in differential recognition of the DNA damage by cellular
proteins. In such a case the conformational changes between the mono- and bi-functional adduct
are so different that the hydrogen bond may not have the same function; however, the details of
protein recognition of Pt-DNA adducts are poorly understood, and this feature may be important
in an as yet unknown role. It is intriguing that the SS-isomer of oxaliplatin, which is inactive, is
unable to form this hydrogen bond. This evidence suggests that the interaction is important, even
if the mechanism behind it is not yet clear.
Second, the platinum-DNA adduct causes similar distortion of the base pair step on the 5'
side of the lesion, regardless of the mono- or bidentate nature of binding, which moves the base
pair containing the platinum adduct out into the major groove (see Fig. 2.9). These changes are
not as dramatic for the cDPCP mono-adduct compared to the cisplatin intrastrand cross-link.
Again, the consequences of these structural changes in terms of cellular recognition and
cytotoxicity have not been determined, but it is compelling that pyriplatin binding cause similar
distortions to DNA geometry as cisplatin while forming a fundamentally different type of adduct.
I G7*
C18
Figure 2.9. The platinated base pair (blue) overlaid with ideal B-form DNA (gray). Platinum
binding forces the DNA bases out into the major groove, causing significant increases in the shift
and slide values of the base pair step to the 5' side of the adduct.
When discussing the mechanism of cytotoxicity of pyriplatin in cancer cells, the
possibility must be entertained that in vivo, the heterocyclic amine is labilized and the adduct
closes to form a bifunctional cross-link. However, all evidence points to the contrary. Leng and
coworkers demonstrated that when complexes of the form [Pt(NH 3) 2(Am)Cl]"* bind to double
stranded DNA, the heterocyclic amine ligand (Am) can be released and bifunctional adducts
form, depending on the nature of the ligand. This reaction occurs most readily when Am = [Nmethyl-2,7-diazapyrenium]*; 37 ,38 however, cDPCP adducts were very stable and showed no
degradation to intra- or interstrand cross-links.8 These results, combined with those of
immunochemical studies that demonstrate that cisplatin and pyriplatin form different adducts,
and replication mapping experiments showing inhibition of DNA polymerases at single dG
sites, 7 strongly suggest that monofunctional adducts of pyriplatin persist in tumors and are
ultimately responsible for the activity of the compound.
Despite the similarity between cisplatin- and pyriplatin-DNA adducts, the global
structures of each nucleic acid double helix are significantly different. These results suggest that
the two platinum antitumor compounds may inhibit transcription and destroy cancer cells
through different mechanisms. Compared to cisplatin, pyriplatin causes only minor distortions to
double helical DNA upon binding to a guanine N7 atom in the major groove. Characteristics of
the cisplatin 1,2-intrastrand d(GpG) cross-link include a dramatic roll angle between the bound
guanines and a 400 bend towards the major groove.' 4 These structural distortions may cause a
decrease in transcription levels, which triggers either nucleotide excision repair or apoptosis.
Evidence from a recent X-ray crystal structure of an RNA polymerase II elongation complex
containing a cisplatin 1,2-d(GpG) intrastrand cross-link on the DNA template strand indicates
that the platinum adduct inhibits transcription by prohibiting translocation of the cross-link from
the +2/+3 site to the +2/+1 site (the +1 site is where ribonucleotide incorporation occurs. This
barrier stems from the inability of the covalently linked dinucleotide to twist by ~904 for
crossing a protein bridge helix near the +2/+1 site.3 6 This finding provides a structural clue for
understanding transcription inhibition by cisplatin intrastrand cross-links.
The model of cDPCP-damaged DNA suggests a different mechanistic hypothesis for
pyriplatin inhibition of transcription by pol II. Monofunctional adducts of pyriplatin would not
be subjected to the same translocation barrier crossing into the +1 active site because cDPCP
2
binds covalently only to a single DNA base. Modeling of the cis-{Pt(NH 3)2(py)} -dG adduct
into the template strand of the elongation complex suggested the absence of a significant barrier
to translocation between the +2 and +1 sites. However, when the adduct is modeled in the +1
site, where the incoming NTP is matched to the template strand, of an elongation complex solved
crystallographically, 39 it appears that the pyridine ligand would sterically clash with the bridge
helix, in effect twisting the base out of its native conformation (depicted in Fig. 2.10.a)). This
distortion could lead to NTP misincorporation or inability to close the trigger loop that is
required for catalysis, which in turn would stall the polymerase. This hypothesis explains why
pyriplatin damage inhibits transcription, whereas adducts of the trans isomer and other
monofunctional platinum compounds like [Pt(dien)Cl]Cl, which would not cause such steric
hindrance, are much less potent inhibitors. Transcription is strongly inhibited by pyriplatin both
in cell extracts and in live cells, but a significantly higher level of platination with [Pt(dien)Cl]*
is required to block transcription to the same extent.
New structural evidence has recently been obtained that reveals an alternative possibility
for pyriplatin inhibition of transcription.40 The yeast RNA polymerase 1I elongation complex was
crystallized with a cis-{Pt(NH 3)2(py)} 2+-dG adduct in the +1 site, both before and after ligation
of CMP onto the RNA strand. In the pre-elongation state with an empty NTP addition site, the Pt
lesion adopts a conformation similar to the one modeled in Fig. 2.10.a, with the aromatic ring
pointed towards the 5' end of the DNA template and lying near the bridge helix. However, the
guanine base position still accommodates the incoming CTP. After RNA elongation the complex
is inhibited in the pre-translocation state, meaning that the Pt-dG lesion stalls in the +1 site and
cannot move upstream. In this post-elongation, pre-translocation state (shown Fig. 2.10.b), the
cis-{Pt(NH 3)2(py)} 2+-dG conformation changes dramatically; the pyridine ring is directed
toward the 3' end of the DNA template and the Pt ammine ligand trans to the pyridine ring
interacts with the bridge helix through hydrogen bonding to a threonine residue that is conserved
from yeast to human pol II. This interaction may be responsible for stabilizing the elongation
complex and preventing translocation and subsequent transcription.
(A)
RNA
Template DNA
Non-template DNA
(B)
RNA
Template DNA
A828
Non-template DNA
Figure 2.10. (on previous page) (A) Active site of RNA polymerase II, with a cDPCP-dG
adduct (shown in orange) modeled into template DNA (blue) at the +1 site, where incoming
NTPs are matched and added to synthesized RNA (red). Complementary, non-template DNA is
shown in green. This model demonstrates how cDPCP adducts may shift the template base out of
its native conformation (shown in dark blue) by steric interactions between the pyridine ligand
and the Pol 1I bridge helix, shown in gray. Inset: Space-filling views of the Pt adduct and bridge
helix. Pol II coordinates: PDB code 2NVQ, cDPCP-dG coordinates: this work. (B) Recently
published X-ray crystal structure of cDPCP-dG adduct in the +1 site of pol IL.CMP is added to
the RNA strand but the complex stalls in the pre-translocation state. Hydrogen-bonding
interactions between a Pt ammine ligand and the bridge helix may contribute to this inhibition
mechanism.
Conclusions
A DNA dodecamer duplex containing a site-specific cis-{Pt(NH 3)2(pyridine)} 2 +-dG
adduct was synthesized for X-ray diffraction studies. The result is the first structure of a
biologically active monofunctional platinum(II) compound covalently bound to DNA. In contrast
to cisplatin-damaged DNA, which is bent by ~40' toward the major groove, the dodecamer
containing a cis-{Pt(NH 3 )2 (pyridine)} 2+-dG lesion remains linear, without significant global
distortion. However, several characteristics of the platinum-DNA adduct are shared with
bifunctional Pt-DNA cross-links. A hydrogen bonding interaction between the ammine ligand cis
to the guanine and the carbonyl oxygen atom 06 of the binding nucleobase is analogous to a
similar feature observed in oxaliplatin-DNA adducts, and this interaction has been previously
implicated to impart cytotoxicity to the compound. Also in this model is a distorted base pair
step to the 5' side of the platinum lesion, marked by unusually large shift and slide values, which
is shared by bifunctional cisplatin cross-links. Because of its selective uptake by OCT1, the
monofunctional platinum(II) cation cDPCP has potential to enter pre-clinical trials as a treatment
for colorectal cancer with minimal toxicity to healthy tissue. This complex would be added to the
list of "rule-breakers" that do not follow the classical structure-activity relationship for platinum
antitumor compounds, but display activity against tumors.
References
(1)
Jamieson, E. R.; Lippard, S. J. Chem. Rev. 1999, 99, 2467-2498.
(2)
Wang, D.; Lippard, S. J. Nat. Rev. Drug Discov. 2005, 4, 307-320.
(3)
Cleare, M. J.; Hoeschele, J. D. Bioinorg. Chem. 1973, 2, 187-2 10.
(4)
Farrell, N. In Metal Ions in BiologicalSystems; Sigel, A., Sigel, H., Eds.; Marcel Dekker:
New York, 2004; Vol. 42, p 251-296.
(5)
Perez, J. M.; Fuertes, M. A.; Alonso, C.; Navarro-Ranninger, C. Crit. Rev. Oncol.
Hematol. 2000, 35, 109-120.
(6)
Hollis, L. S.; Amundsen, A. R.; Stem, E. W. J Med. Chem. 1989, 32, 128-136.
(7)
Hollis, L. S.; Sundquist, W. I.; Burstyn, J. N.; Heiger-Bemays, W. J.; Bellon, S. F.;
Ahmed, K. J.; Amundsen, A. R.; Stem, E. W.; Lippard, S. J. Cancer Res. 1991, 51, 18661875.
(8)
Payet, D.; Leng, M. In StructuralBiology: The State of the Art; Sarma, R. H., Sarma, M.
H., Eds.; Adenine Press: Albany, 1994; Vol. 2, p 325-333.
(9)
Zhang, S.; Lovejoy, K. S.; Shima, J. E.; Lagpacan, L. L.; Shu, Y.; Lapuk, A.; Chen, Y.;
Komori, T.; Gray, J. W.; Chen, X.; Lippard, S. J.; Giacomini, K. M. Cancer Res. 2006,
66, 8847-8857.
(10)
Lee, K.-B.; Wang, D.; Lippard, S. J.; Sharp, P. A. Proc. Natl. Acad. Sci. US.A. 2002, 99,
4239-4244.
(11)
Tremeau-Bravard, A.; Riedl, T.; Egly, J.-M.; Dahmus, M. E. J. Biol. Chem. 2004, 279,
7751-7759.
(12)
Lovejoy, K. S.; Todd, R. C.; Zhang, S.; McCormick, M. S.; D'Aquino, J. A.; Reardon, J.
T.; Sancar, A.; Giacomini, K. M.; Lippard, S. J. Proc.Natl. Acad. Sci. US.A. 2008, 105,
8902-8907.
(13)
Marzilli, L. G.; Saad, J. S.; Kuklenyik, Z.; Keating, K. A.; Xu, Y. J. Am. Chem. Soc.
2001, 123, 2764-2770.
(14)
Takahara, P. M.; Frederick, C. A.; Lippard, S. J. J. Am. Chem. Soc. 1996, 118, 1230912321.
(15)
Todd, R. C.; Lippard, S. J. In Platinum and Other Heavy Metal Compounds: Molecular
Mechanisms and ClinicalApplications;Bonetti, A., Howell, S. B., Leone, R., Muggia, F.,
Eds.; Humana Press: Totowa, 2009, p 67-72.
(16)
Spingler, B.; Whittington, D. A.; Lippard, S. J. Inorg. Chem. 2001, 40, 5596-5602.
(17)
Silverman, A. P.; Bu, W.; Cohen, S. M.; Lippard, S. J. J Biol. Chem. 2002, 277, 4974349749.
(18)
Dhara, S. C. Indian J. Chem. 1970, 8, 193-194.
(19)
Caruthers, M. H. Acc. Chem. Res. 1991, 24, 278-284.
(20)
Drenth, J. Principles ofProteinX-ray Crystallography; Springer: New York, 1994.
(21)
Ducruix, A.; Giege, R. Crystallization of Nucleic Acids and Proteins: A Practical
Approach; Oxford University Press: New York, 1999; Vol. 2.
(22)
Scott, W. G.; Finch, J. T.; Grenfell, R.; Fogg, J.; Smith, T.; Gait, M. J.; Klug, A. J Mol.
Biol. 1995, 250, 327-332.
(23)
Otwinowski, Z.; Minor, W. Methods Enzymol. 1997, 276, 307-326.
(24)
Fortelle, E. d. L.; Bricogne, G. Methods Enzymol. 1997, 276, 472-494.
(25)
Emsley, P.; Cowtan, K. Acta Crystallogr.2004, D60, 2126-2132.
(26)
Murshudov, G. N.; Vagin, A. A.; Dodson, E. J. Acta Crystallogr. 1997, D53, 240-255.
(27)
Winn, M. D.; Isupov, M. N.; Murshudov, G. N. Acta Crystallogr.2001, D57, 122-133.
(28)
Painter, J.; Merritt, E. A. Acta Crystallogr.2006, D62, 439-450.
(29)
Painter, J.; Merritt, E. A. J. Appl. Crystallogr.2006, 39, 109-111.
(30)
BrUnger, A. T.; Adams, P. D.; Clore, G. M.; DeLano, W. L.; Gros, P.; Grosse-Kunstleve,
R. W.; Jiang, J.-S.; Kuszewski, J.; Nilges, M.; Pannu, N. S.; Read, R. J.; Rice, L. M.;
Simonson, T.; Warren, G. L. Acta Crystallogr.1998, D54, 905-92 1.
(31)
Lu, X.-J.; Olson, W. K. Nucleic Acids Res. 2003, 31, 5108-5121.
(32)
Comess, K. M.; Costello, C. E.; Lippard, S. J. Biochemistry 1990, 29, 2102-2110.
(33)
Bauer, C.; Peleg-Shulman, T.; Gibson, D.; Wang, A. H.-J. Eur. J Biochem. 1998, 256,
253-260.
(34)
Wu, Y.; Bhattacharyya, D.; King, C. L.; Baskerville-Abraham, I.; Huh, S.-H.; Boysen,
G.; Swenberg, J. A.; Temple, B.; Campbell, S. L.; Chaney, S. G. Biochemistry 2007, 46,
6477-6487.
(35)
Sharma, S.; Gong, P.; Temple, B.; Bhattacharyya, D.; Dokholyan, N. V.; Chaney, S. G. J
Mol. Biol. 2007, 373, 1123-1140.
(36)
Damsma, G. E.; Alt, A.; Brueckner, F.; Carell, T.; Cramer, P. Nat. Struct. Mol. Biol.
2007, 14, 1127-1133.
(37)
Malinge, J.-M.; Sip, M.; Blacker, A. J.; Lehn, J.-M.; Leng, M. Nucleic Acids Res. 1990,
18, 3887-3891.
(38)
Anin, M.-F.; Gaucheron, F.; Leng, M. Nucleic Acids Res. 1992, 20, 4825-4830.
(39)
Wang, D.; Bushnell, D. A.; Westover, K. D.; Kaplan, C. D.; Komberg, R. D. Cell 2006,
127, 941-954.
(40)
Wang, D.; Zhu, G.; Huang, X.; Lippard, S. J. Proc. Natl. Acad. Sci. U.S.A. 2010, 107, in
press.
Chapter 3. Reactions between Phosphate and Carbonate Complexes of Cisplatin with
Nucleic Acids: Investigations of the Resulting Pt-DNA Adduct Structure and Yield
Research in this chapter has been published in J Am. Chem. Soc. 2007, 129 (20), 6370-6371
Introduction
Activation of the anticancer drug cisplatin, cis-diamminedichloroplatinum(II), for binding
to biological targets involves replacement of chloride ligands with water molecules.' Aquated
cisplatin can bind to purine bases on DNA, forming intrastrand cross-links that destroy cancer
cells.2 The intermediacy in this process of platinum complexes with other biologically abundant
anions such as acetate, 3 phosphate, 3 and thiolate, 4 has been explored in depth. Acetate,
phosphate, and pyrophosphate anions all increase the extent of chloride ion release from cisplatin
in solution and decrease the rate of cisplatin binding to nucleotide targets.3 Other data suggest
that interactions between cisplatin and the phosphodiester backbone of DNA may serve as
intermediates in binding to nucleobases.5 Platinum binding by intracellular thiols such as
glutathione are well documented 4 and postulated to play a major role in deactivating platinum
drugs, reducing their antitumor efficacy.6
Recent work draws attention to the reaction of cisplatin with carbonate ion and proposes
a potential role of platinum carbonato species in mediating cellular uptake, DNA binding, and
anti-tumor properties.7-9 Because these species are formed after introducing cisplatin into the
bloodstream during chemotherapy, it is important to evaluate their role in binding to nuclear
DNA and contribution to triggering cell death. In this chapter the effects of phosphate and
carbonate on cisplatin binding to single-stranded DNA in vitro are reported, to determine the
effects of these complexes on the yield and structure of resulting Pt-DNA adducts. The results,
which are fully consistent with early immunochemical studies of platinum anticancer drugs,' 0 ' 2
demonstrate that at physiological concentrations of carbonate the major DNA adducts are
bifunctional cross-links. Like phosphate ion and other biological nucleophiles, the chief
consequence of carbonate binding is to reduce the amount of platinum available for DNA
modification.
Experimental
Materials. Potassium tetrachloroplatinate(II) used to synthesize cisplatin13 was obtained as a gift
from Engelhard Corporation (now BASF, Iselin, NJ). Phosphoramidites, columns, and other
reagents for oligonucleotide synthesis were purchased from Glen Research. All other reagents
and solvents were obtained from commercial sources and used without further purification. Calf
intestinal phosphatase was purchased from New England Biolabs, and nuclease P1 from MP
Biomedical. Oligonucleotide synthesis was performed with an Applied Biosystems 392
DNA/RNA automated synthesizer. Analytical and preparative HPLC separations were performed
with an Agilent 1200 HPLC equipped with an automated fraction collector. Atomic absorption
and UV-Vis spectroscopy were performed using a Perkin-Elmer AAnalyst 300 system and a HP
8453 UV-visible spectrophotometer, respectively. HPLC-coupled electrospray ionization mass
spectrometry (ESI-MS) was performed on an Agilent 1100 series HPLC with an MSD trap.
Reaction of cisplatin with a single-stranded DNA. A 14 base synthetic oligonucleotide having
sequence 5'-d(TTCACCGGAATTCC)-3' was allowed to react with cisplatin (1.2 equiv) in 24
mM carbonate, phosphate, or HEPES buffer, at either pH 6.8 or 7.4, and in the presence of 5 mM
NaCl in O-ring sealed Eppendorf tubes. Cisplatin concentrations investigated were 10, 20, 40,
100, and 200 gM. Cisplatin solutions were freshly prepared at a concentration of 3 mM in 41
mM NaCl and used immediately. All reactions were incubated at 37 'C for 24 h in the dark and
then analyzed by ion-exchange HPLC. Separation was accomplished on a 6.2 x 80 mm Agilent
Zorbax Oligo column (see method RCT.004A, Appendix C) To isolate products, peak fractions
at 27.0, 28.5, and 45.4 min were collected with an automated fraction collector for characterization (see Fig. 3.1). Combined fractions were dialyzed against water in the dark at 4 'C,
lyophilized to dryness, and desalted with Waters Sep-pak C18 cartridges. For reactions under
inert nitrogen atmosphere, a solution of 41 mM NaCl (4 mL) was degassed by 4 freeze-pumpthaw cycles, then transferred under nitrogen to a vial containing 1.24 mg (4.1 mmol) of cisplatin,
to make a 1 mM stock solution. In a separate vial, a solution of 14mer (7.5 nmol) in 890 pL of 5
mM NaCl, 24 mM HEPES pH 7.4 was degassed by 4 freeze-pump-thaw cycles. To this solution
was added 9 pL of cisplatin solution (9 nmol) by microsyringe under N2. The solution was
incubated at 37 'C for 24 h and analyzed as before. All yields are reported relative to the reaction
in HEPES buffer, which remains constant in all samples.
Mass spectrometry. ESI mass spectra were collected on an Agilent 1100 SL MSD trap in
negative ion mode. The oligonucleotides were desalted on-line by HPLC. Samples were passed
through an Agilent Extend C18 2.1 x 150 mm, 3.5 pm column using method RCT.010M (in
Appendix C). The mass spectrum of the major platinated product is shown in Fig. 3.2.
Enzymatic digestion. The procedure for digestion of oligonucleotides to break down DNA
products to the nucleoside level was modified from that described previously.' 4 In 100 pL of
digestion buffer (1 mM ZnCl 2, 20 mM Na acetate pH 5.2), each of the two collected products
and the unplatinated 14mer (5 nmol each) were mixed with 10 ptL (10 U) of nuclease Pl. The
samples were incubated at 37 'C overnight. To each sample was added 5 tL of 1.5 M Tris-HCl,
pH 8.8, and 1 pL (10 mU) of calf intestinal phosphatase (CIP). After 4 h of incubation at 37 'C,
6 ptL of 0.1 N HCl was added to precipitate the protein, and the samples were vortexed and
centrifuged at 13,500 RPM for 5 min. Each sample was analyzed by LC-MS on a Supelcosil LC18-S, 2.1 x 250 mm, 5 pm column using method RCT.009M in Appendix C. Peaks areas were
normalized to account for differences in extinction coefficients at 260 nm, and then peak ratios
were calculated relative to the thymine peak. Thymine is the least reactive base toward cisplatin,
so the peak area should remain constant upon DNA platination.
160.
140-
{
-Carbonate
120
-
,
Phosphate
HEPES
80
<6040
2
20
0
10
20
30
40
50
60
70
80
Time (ai)
Figure 3.1. HPLC chromatograms of the reaction of 40 ptM cisplatin with the oligonucleotide,
5'-TTC-ACCGGAATTCC-3', in 24 mM carbonate, phosphate, or HEPES buffer, pH 7.4, and 5
mM NaCl. The major product (peak 1) elutes at 27.0 min, and a minor peak (peak 2) appears at
28.5 min. The unplatinated oligonucleotide elutes at 45.4 min.
Results
Product characterization. The reaction of cisplatin with single stranded DNA was investigated
in several buffer systems to investigate the effect of carbonate or phosphate ion on cisplatinDNA binding. Two major products, denoted 1 and 2 in Figure 3.1, were isolated from all
platination reactions by HPLC with retention times of 27.0 and 28.5 min, respectively. Unreacted
starting material was also collected at 45 min. The mass of 1, as revealed in the negative ion ESI
spectrum, is 4424.9 ± 1.7 Da (Fig. S4), in agreement with the expected value of the 14mer
bearing a {Pt(NH 3)2} 2+ lesion (4425.8 Da). The Pt/DNA ratio was 1.09 ± 0.04. Product 2 has a
determined mass of 4424 ± 2 Da and a Pt/DNA ratio of 1.18 ± 0.03. These results indicate that
both products contain bifunctional cross-links, because monofunctional Pt-DNA adducts would
have higher mass arising from the presence of a third Pt ligand.
-4
.7
Figure 3.2. ESI (-) spectrum of major product peak 1. 631.5 m/z = [M - 7H]7 ~,736.3 m/z = [M 6H] 6-, 883.8 m/z = [M - 5H] 5 , 1105.5 m/z = [M - 4H] 4-. Found mass = 4424.9 ± 1.7 Da.
Calculated mass of 14mer with {Pt(NH 3)2}2+ adduct = 4425.8 Da.
Enzymatic cleavage with nuclease P1 and calf intestinal phosphatase breaks down DNA
to its nucleoside components, and the ratios of each base can be quantitated by HPLC using the
relative UV absorbance ratios of each nucleobase. Digestion of the unmodified strand yields the
expected ratio of each deoxynucleoside. Digestion of 1 (Fig. 3.3) revealed that the free dG peak
disappears nearly completely, indicating that both guanines are bound by Pt, and a new peak
grows in (marked by *). The negative ion mass spectrum of this peak (Fig. 3.4) contained signals
at 884.5 m/z, 901.9 m/z, 938.8 m/z, and 960.8 m/z; these signals were assigned to [{Pt(NH3)2}2+_
d(GpGp)-OH]~, [{Pt(NH 3)2 }2+-d(GpGp)-H]~, [{Pt(NH 3)2 }2+-d(GpGp)+Cl]~, and [{Pt(NH 3)2 }2+_
d(GpGp)+AcO]-, respectively, allowing this new digestion peak to be identified as the 1,2{Pt(NH 3)2 }2+-d(GpG) cross-link. All of these peaks contain isotope patterns consistent with
platinum-containing species. Minor peaks in the mass spectrum were assigned to the 1,2-
{Pt(NH 3)2 }2 -d(GpA) lesion. Because the adenosine peak area is only reduced by 7% compared
to the unplatinated oligomer, this lesion is a very minor component of the sample. Although 1,2(ApG) crosslinks can account for ~20% of all platinum-DNA adducts, 1,2-(GpA) adducts are not
observed in vivo.1 5
unplatinated 14mer
2SOO
dG T
dC
ICale.
2000
dC
dG
T
S1500
5
2 2.2
4 4.0
dAI 3
1000
Obs.
49
3.0
-
500 -
10
20
40
30
Time (mn)
2000
peak 1
18001600 -
Calc, Obs.
1400
ldCI5
4.81
1200 -
dG
0.1
T
1000-
2
4 4.0
SOO
600
400
200
0
''
Time (min)
peak 2
1800
1600
Catc. Obs.
1400
Ido 5 4.3
dG 2 0.9
T 4 4.0
1200
1000
dA 3 2.4
800
600
400
200
0
I -Ir
20
Time (min)
Figure 3.3. HPLC chromatograms of enzymatic digestion of 14mer DNA (top), the major
platinated product (middle) and the minor Pt species (bottom). Peaks arising from platinum
adducts are marked by * and # for the major and minor products, respectively.
Digestion of product 2 (Fig. 3.3) reveals reduction but not complete disappearance of the
free dG peak, as well as growth of a smaller peak at 19.3 min (marked by #) with ESI-MS
signals (Fig. 3.4) at 573.5 m/z, 110.9 m/z, 1147.8 m/z, and 1169.8 m/z. These m/z ratios were
assigned as [{Pt(NH 3)2}2+-d(CpGpG)+Cl] 2-, [{Pt(NH 3)2 }2+-d(CpGpG)-H],
[{Pt(NH3)2}2 +_
d(CpGpG)+Cl]-, and [{Pt(NH 3)2}2+-d(CpGpG)+AcO]~, respectively. Analysis of the peak ratios
of the digest reveals that the deoxycytidine, deoxyguanosine, and deoxyadenosine peaks were all
reduced relative to the starting material. The dC peak was reduced by 12% compared to the
unplatinated oligo, and the dA peak was reduced by 20%. The dG peak was reduced by 58%.
The identity of the 1,3-{Pt(NH 3)2}2+-d(CpGpG) cross-link was later confirmed by synthesizing a
standard; LC/MS analysis showed a peak with the same retention time and mass spectrum.
*
.O,
#Lil
w~ane
MyG
9306
92031
s-
ow
O60
amo
wg
0
an0
0
no9
0lo
4
1M
0
"D
O
160
f
wis
Fig. 3.4. ESI (-) mass spectra of Pt-DNA cross-links arising from enzymatic digestion of singlestranded DNA. Peak assignments are given in the text.
Yield of Pt-DNA adducts. As shown by the chromatograms of each reaction (Fig 3.1), the
product profile of the reaction of cisplatin and a 14mer oligonucleotide does not change,
regardless of whether the reaction occurs in HEPES, phosphate, or carbonate buffer. Two major
products, denoted peaks 1 and 2, are formed in all reactions. In both carbonate and phosphate
buffers, however, the reaction is significantly inhibited relative to the reaction in HEPES buffer,
particularly at concentrations of cisplatin below 40 mM.
NO%
60%
20 %
0
50
(K
150
200
(Cisplatin]1, pM
Figure 3.5. Yield of platinated 14mer ssDNA in 24 mM carbonate (black, pH 7.4, and blue, pH
6.8) or phosphate (green, pH 7.4, or red, pH 6.8) buffer, normalized to the yield in HEPES
buffer, as a function of cisplatin concentration.The presence of both carbonate and phosphate
ions strongly inhibits the reaction at low concentrations of cisplatin.
Figure 3.5 show a comparison of the yields of each product as a function of cisplatin
concentration in each buffer system at pH 7.4. At pH 6.8, a similar trend is observed; cisplatin
binding to DNA is largely inhibited in carbonate and phosphate buffers relative to its binding
capacity in HEPES buffer. Whereas the yield in HEPES buffer does not change by lowering the
pH, the yield of platinated oligo in carbonate and phosphate buffer increases slightly relative to
the yield at pH 7.4. At both pH 6.8 and 7.4, the 1,3-intrastrand cross-link accounts for a slightly
higher percentage of total adducts in phosphate- or carbonate-containing samples compared to
that for reactions in HEPES buffer (Fig. 3.1, compare relative peak areas of 1 and 2).
Bifunctional cross-links constitute the primary binding mode of cisplatin to DNA in all buffer
systems, but the relative amounts of 1,2- and 1,3- adducts show a dependence on the platinum
leaving group.
Because all aqueous solutions contain dissolved C0 2, they also have some amount of
carbonate ion. Carbonate was completely removed in some of the HEPES samples by degassing
the solutions prior to reaction. In this way, the binding of cisplatin to DNA could be investigated
in a carbonate-free environment. The results of these experiments were the same as those in
non-degassed HEPES buffer, indicating that the residual amount of carbonate ion in these
samples does not significantly affect the reaction.
Discussion
The
reaction
of cisplatin
with
a single-stranded
14mer
oligonucleotide,
5'-
TTCACCGGAATTCC-3', was investigated in 24 mM carbonate, phosphate, and HEPES buffers
at pH 6.8 and 7.4, containing 5 mM NaCl. The DNA contains a single reactive GG site, allowing
for both mono- and bifunctional platinum adducts to fonn. Cisplatin concentrations from 10 to
200 pM were investigated, and the DNA concentration was adjusted to maintain a fixed Pt:DNA
ratio of 1.2:1. After 24 h of incubation in the dark at 37 'C, reaction products were purified by
ion exchange high performance liquid chromatography and characterized by UV and AA
spectroscopy, enzymatic digestion and ESI-MS.
In all buffer systems, the major product of the reaction was determined to be the 1,2{Pt(NH 3)2 }2+-d(GpG) cross-link. The mass of this product, as revealed in the negative ion ESI
spectrum, is 4424.9 ± 1.7 Da, in agreement with the expected value of the 14mer bearing a
{Pt(NH 3)2 }2+ lesion (4425.8 Da). After enzymatic digestion of the products with nuclease P1 and
CIP and separation of the resulting deoxynucleosides by LC-MS, the 1,2-{Pt(NH 3)2}2+-d(GpG)
adduct was directly observed (Fig. 3.4). A minor product detected by the same methods was a
DNA strand containing a 1,3-{Pt(NH 3)2}2 +-d(CpGpG) cross-link. Cytosine is not very reactive
toward cisplatin, although binding of platinum to the cytosine N3 position has been observed in
an oligonucleotide.16
Identification of the 1,3-intrastrand cross-link initially was ambiguous from the nuclease
digestion data. ESI-MS of the product and Pt/DNA ratios clearly assign 2 as a bifunctional crosslink. However, because the 1,3-{Pt(NH 3)2}2+-d(CpGpG) moiety was observed in the digestion,
the free dG peak should have disappeared if the product were 100% pure. Instead, ~0.9
guanosine per DNA was observed in the chromatogram. This observation may be the result of
incomplete removal of the internal dG in the cross-link, where some of the bridging base was
removed, leading to the presence of an unmodified dG peak, with some of the 1,3-{Pt(NH 3)2}2+_
d(CpGpG) remaining intact and eluting as a new peak. No signal for {Pt(NH 3)2(dC)(dG)} 2+was
observed in the digestion results however. Another possibility is that some fraction of the
product is a 1,3-{Pt(NH 3)2}2+-d(CpCpG) cross-link. These results indicate that the second
collected fraction 2 may not be a single product; however, verification of the 1,3-{Pt(NH 3)2} 2+_
d(CpGpG) cross-link by preparation of a synthetic standard clearly demonstrates that this
product constitutes a significant fraction of the product, and it is possible that other minor
components of the sample have the same structural composition, a 1,3-{Pt(NH 3)2} 2+ intrastrand
cross-link, that would lead to identical retention times on an HPLC column.
A higher percentage of 1,3-intrastrand cross-links was observed in carbonate and
phosphate buffers relative to the reaction in HEPES buffer, as indicated by the relative peak
areas of 1 and 2 from chromatograms in Fig. 3.1. This observation may indicate that platinum
carbonato or phosphato complexes go through different reaction intermediates than when
chloride ion is the leaving group. Indeed, the product profile of cisplatin and carboplatin are
markedly different both in vitro and in vivo; cisplatin binding to DNA produces -90% 1,2-
intrastrand cross-links, 0 1 2 whereas the major product of carboplatin binding, ~35% of total
adducts, is the 1,3-d(GNG) cross-link.17 The only difference between the two molecules is the
leaving group, two chloride ions for cisplatin or cyclobutane-dicarboxylate in the case of
carboplatin. The different reaction intermediates may be a result of varying kinetics for
replacement of the leaving groups, the difference of one bidentate vs. two monodentate leaving
groups, or a combination of both factors.
The product profile of the reaction of cisplatin and the 14mer oligonucleotide is
unaffected by the choice of HEPES, phosphate, or carbonate buffer, in that bifunctional
platinum-DNA cross-links are the dominant products of all three reactions. In both carbonate and
phosphate buffers, however, the reaction is significantly inhibited relative to the reaction in
HEPES buffer, particularly at concentrations of cisplatin < 40 mM.
At pH 6.8, the yield of platinated 14mer in carbonate and phosphate buffer increases
slightly relative to the yield at pH 7.4. The yield of platinated DNA in HEPES buffer is
unaffected by pH. The pKa of carbonate (for the equilibrium CO 2 + H2 0
and the second pKa of phosphate (H2 PO4-
HP0
2
4
I
HCO3-+ H*) is 6.1,18
-I+ H*) is 7.09. Therefore at lower pH, a
larger fraction of both carbonate and phosphate ligands will be protonated, making them better
leaving groups. This pH dependence provides strong evidence that inhibition by these anions is
due to complexation with the platinum(II) center.
Aquated cisplatin complexes are cations; binding to DNA, a polyanion, is thus assisted
by electrostatic attractions. Cisplatin carbonato or phosphato complexes, however, are either
neutral or anionic species, depending on the binding mode and protonation state of the ligand,
which have not been conclusively established. The altered charge may therefore explain why
carbonate and phosphate inhibit binding to DNA. Carbonate is a better ligand for platinum than
phosphate,19 which renders it a stronger inhibitor of the reaction.
The data presented here show that the presence of physiological concentrations of carbonate and phosphate inhibit the binding of cisplatin to single-stranded oligonucleotides through
formation of cisplatin carbonato and phosphato complexes, respectively. Interactions between
cisplatin and other biological nucleophiles such as glutathione 4 or hydroxide iono "deactivate"
cisplatin through formation of relatively unreactive platinum complexes. The experiments here
suggest that carbonate and phosphate ions may perform a similar function. Physiological
concentrations of phosphate and carbonate in interstitial fluid are approximately 5 mM and 24
mM, respectively, and the concentrations of these species in the cell are approximately 80 mM
and 12 mM, respectively. 2 ' Along with proteins and nucleic acids, these biological anions
provide numerous options for binding cisplatin once it is aquated inside the cell (see Fig. 3.6).
H
HN
H N,
'
Cl-N/
HC1
Pt
\N OH *
HN\ p0H
H4N/
\Ci
(b) Carbonate
(a) Aquation
- -+
- - -
\C1
\OH,
N
0
I
]
HN /\C1
H3N
NNtPt
+HHN
H ,N
H N \ t0CO
2+I
HN/
O
H3N
0
-
HsN\ ptO
,-
H3N
Ct
OPO H
Ct
A
S
OPP
0
HN
H,,N
SMet
H
N
2-
HPN-t
co2
Figure 3.6. Routes for biological processing of cisplatin. Upon entering the cell, cisplatin is
aquated (a). The activated platinum species can then react with carbonate (b) or phosphate (c) in
a mono- or bidentate manner. Reaction with glutathione results in the formation of a bis(glutathionato)-platinum complex (d). 4 Platinum can also bind sulfur-containing amino acids,
such as Met in human albumin (e). 2 Finally, cisplatin can react with purine bases of both DNA
and RNA, forming intra- and interstrand cross-links (f).
Conclusions
The effects of biological anions on the yield and structure of cisplatin-DNA adduct
formation on single-stranded DNA were investigated using HPLC, enzymatic digestion, and
mass spectrometry. Carbonate and phosphate ions form complexes with cisplatin that inhibit
binding to nucleic acids. The results also demonstrate that cisplatin binds to DNA in a
bifunctional manner in the presence of both coordinating (carbonate, phosphate) and weakly
coordinating (HEPES)
buffers.
Cisplatin forms
primarily
bifunctional
adducts under
physiologically relevant conditions. Intrastrand cross-links between adjacent guanines have
previously been assigned as the primary cisplatin-DNA adduct in experiments conducted both in
vitro and in vivo with immunochemical detection, accounting for 65% of total platinum-DNA
lesions. 10 - 2 The present findings demonstrate that the presence of carbonate or phosphate does
not affect the preference of cisplatin to form such intrastrand adducts.
References
(1)
Bancroft, D. P.; Lepre, C. A.; Lippard, S. J. J Am. Chem. Soc. 1990, 112, 6860-6871.
(2)
Jamieson, E. R.; Lippard, S. J. Chem. Rev. 1999, 99, 2467-2498.
(3)
Segal, E.; Pecq, J.-B. L. Cancer Res. 1985, 45, 492-498.
(4)
Ishikawa, T.; Ali-Osman, F. J Biol. Chem. 1993, 268, 20116-20125.
(5)
Campbell, M. A.; Miller, P. A. Biochemistry 2008, 47, 12931-12938.
(6)
Kartalou, M.; Essigmann, J. M. Mutation Res. 2001, 478, 23-43.
(7)
Centerwall, C. R.; Goodisman, J.; Kerwood, D. J.; Dabrowiak, J. C. J. Am. Chem. Soc.
2005, 127.
(8)
Binter, A.; Goodisman, J.; Dabrowiak, J. C. J. Inorg. Biochem. 2006, 100, 1216-1224.
(9)
Di Pasqua, A. J.; Goodisman, J.; Kerwood, D. J.; Toms, B. B.; Dubowy, R. L.;
Dabrowiak, J. C. Chem. Res. Toxicol. 2007, 20, 896-904.
(10)
Terheggen, P. M. A. B.; Floot, B. G. J.; Scherer, E.; Begg, A. C.; Fichtinger-Schepman,
A. M. J.; den Engelse, L. Cancer Res. 1987, 47, 6719-6725.
(11)
Fichtinger-Schepman, A. M. J.; Oosterom, A. T. v.; Lohman, P. H. M.; Berends, F.
CancerRes. 1987, 47, 3000-3004.
(12)
Fichtinger-Schepman, A. M. J.; Van der Veer, J. L.; Den Hartog, J. H. J.; Lohman, P. H.
M.; Reedijk, J. Biochemistry 1985, 24, 707-713.
(13)
Dhara, S. C. Indian J. Chem. 1970, 8, 193-194.
(14)
Huang, X.; Powell, J.; Mooney, L. A.; Li, C.; Frenkel, K. Free Rad Biol. Med 2001, 31,
1341-1351.
(15)
Gupta, R.; Beck, J. L.; Sheil, M. M.; Ralph, S. F. J Inorg. Biochem. 2005, 99, 552-559.
(16)
Comess, K. M.; Costello, C. E.; Lippard, S. J. Biochemistry 1990, 29, 2102-2110.
(17)
Blommaert, F. A.; van Dijk-Knijnenburg, H. C. M.; Dijt, F. J.; den Engelse, L.; Baan, R.
A.; Berends, F.; Fichtinger-Schepman, A. M. J. Biochemistry 1995, 34, 8474-8480.
(18)
Gross, E.; Kurtz, I. Am. J Phys. Ren. Phys. 2002,283, F876-F887.
(19)
Howe-Grant, M. E.; Lippard, S. J. In Metal Ions in Biological Systems; Sigel, H., Ed.;
Marcel Dekker: New York, 1980; Vol. 11, p 63-125.
(20)
Lippert, B. Coord. Chem. Rev. 1999, 182, 263-295.
100
(21)
Wynsberghe, D. V.; Noback, C. R.; Carola, R. Human Anatomy and Physiology; 3rd ed.;
McGraw-Hill: New York, 1995.
(22)
Ivanov, A. I.; Christodoulou, J.; Parkinson, J. A.; Barnham, K. J.; Tucker, A.; Woodrow,
J.; Sadler, P. J. J Biol. Chem. 1998, 273, 14721-14730.
101
Chapter 4. Structure of Duplex DNA Containing the Cisplatin 1,2-{Pt(NH 3)2}2+-d(GpG)
Cross-link at 1.77
A Resolution
This research is published in J. Inorg.Biochem. 2010, in press.
102
Introduction
Platinum-based therapy remains a highly utilized and effective option in the treatment of
many types of cancer.' After cisplatin (cis-diamminedichloroplatinum(II)) was discovered to
have antitumor properties more than 40 years ago,
much research has focused on unraveling
the mode of action of this compound. Nuclear DNA is an important molecular target for
platinum anticancer compounds, which bind purine bases at the N7 position. The resulting PtDNA damage triggers downstream effects including inhibition of replication and transcription,
cell cycle arrest, and attempted repair of the damaged nucleotides. If the cell cannot remove the
damage then it dies by one of several pathways. 4
Revealing the structural details of Pt-DNA adducts represented a significant milestone in
platinum anticancer research. This information has helped to build structure-activity
relationships that underlie transcription inhibition and cell death. The major adduct of cisplatinDNA binding is a 1,2-intrastrand adduct between adjacent guanine bases; a minor percentage of
1,3-intrastrand and interstrand cross-links also form. 4 The 1,2-d(GpG) cisplatin cross-link was
first characterized crystallographically in a DNA dodecamer duplex in 1996 at 2.6-A resolution.
This structure revealed that the Pt adduct induces a global bend of 35-40' in the DNA duplex and
unwinds the double helix by ~25 .5'6 The major groove is compacted and the minor groove
widened and flattened. A-form DNA comprises the nucleic acid to the 5' side of the Pt crosslink, and B-DNA forms to the 3' side of the 1,2-d(GpG) adduct. The roll angle between
platinated guanine bases is 260 and, as a consequence, results in considerable strain being placed
on the Pt-N7 bonds, displacing the Pt atom out of the guanine ring planes by approximately 1 A
each.
103
In this chapter a high resolution, 1.77 A structure of this complex, a dodecamer duplex
having the sequence 5'-CCTCTG*G*TCTCC-3', where the asterisks denote platination sites, is
reported. 7 This significant increase in resolution offers a much-improved view of the electron
density, particularly in the Pt-DNA adduct region. With these new data in hand, the structure of
the cisplatin 1,2-cis-{Pt(NH 3)2}2+-d(GpG) cross-link on DNA is evaluated at a level not
previously attainable. In particular, new information is presented about the overall DNA
geometry, the square-planar platinum coordination environment, the conformation of the 1,2-cis{Pt(NH 3)2}2+-d(GpG) intrastrand cross-link, and the interactions between Pt-DNA molecules and
water and metal ion components of the crystallization buffer. At 1.77-A resolution the
deoxyribose ring conformations can be clearly delineated from electron density maps, providing
useful information about the nucleic acid structure. Better electron density also exists for the
platinum ammine ligands, so that the Pt-N bond distances and angles are more accurately
determined and now match closely the values expected for a square-planar Pt coordination
compound. The previous unexpected observation that the Pt-N7 bonds to each guanine base are
highly strained is re-visited, in order to ascertain the origin of this property. Finally, located in
the current structure are four octahedral [Mg(H 2O) 6 ]2 + ions that interact with the DNA duplex
through their primary hydration spheres; ordered water molecules that hydrogen bond to both the
nucleic acid and the {Pt(NH 3)2} 2+unit are also defined.
Experimental
Materials. Phosphoramidites, columns, and other reagents for solid-phase oligonucleotide
synthesis were purchased from Glen Research. Potassium tetrachloroplatinate(II), which was
used to synthesize cisplatin according to published procedures,8 was a gift from Engelhard
104
(Iselin, NJ, now BASF). Crystallization reagents and supplies were obtained from Hampton
Research. All other reagents were purchased from commercial suppliers and used without further
purification. Oligonucleotides were prepared in-house using an Applied Biosystems Model 392
DNA/RNA Synthesizer. Liquid chromatography was performed with an Agilent 1200 series
HPLC equipped with a temperature-controlled autosampler and automated fraction collector.
Atomic absorption spectroscopy was performed with a Perkin Elmer AAnalyst 300 system. UVVis spectra were collected on a Hewlett-Packard 8453 spectrophotometer.
Preparation of platinated DNA duplex. The oligonucleotide 5'-d(CCTCTGGTCTCC)-3' (1)
and its complement (2) were synthesized on a 2 x 1.0 gmol scale with dimethoxytrityl (DMT)
groups on by standard solid-phase synthetic methods. The strands were purified by reversedphase high-pressure liquid chromatography (HPLC) on an Agilent SB-300, 9.4 x 250 mm
column by method RCT.005S in Appendix C. After lyophilization, DMT groups were removed
in 80% acetic acid for 30 min at room temperature, and the oligonucleotides were precipitated
with isopropanol and desalted with Waters Sep-Pak C18 cartridges. Reaction of 1 with 1.2 equiv
of cisplatin was carried out in 10 mM 4-(2-hydroxyethyl)piperazine-1-ethanesulfonic acid
(HEPES) pH 6.8 buffer for 14 h at 37 'C. The platinated product was purified by ion-exchange
HPLC on a Dionex DNA-Pac PA-100, 9.4 x 250 mm column with method RCT.002S. Pure
fractions were dialyzed against water overnight, then lyophilized and reconstituted in water,
yielding 5'-d(CCTCTG*G*TCTCC)-3' (3). The site-specifically platinated duplex was prepared
by combining equimolar amounts of 2 and 3 in 200 mM LiCl, 100 mM HEPES pH 7.0, and 50
mM MgCl 2 , heating to 70 'C for 10 min, and cooling to 4 'C over 2.5 h. The duplex was purified
by ion exchange HPLC (RCT.003S). The final product was dialyzed, lyophilized, and desalted as
105
described above. HPLC traces of the purification of single- and double-stranded platinated DNA
are shown in Fig. 4.1.
1800
1600
1400
1200
1000
2
800
600
400
200
0
0
5
10
15
20
25
30
35
40
35
40
Time (min)
3
1400 1200
$
1000
S800
600
400
200
0
0
5
10
15
20
25
30
Time (min)
Figure 4.1. HPLC chromatograms of the purification of single- (peak 1, top) and double- (peak
3, bottom) stranded dodecamer DNA modified with a 1,2-cis-{Pt(NH 3)2}2+-d(GpG) cross-link.
Crystallization and X-ray diffraction data collection. Diffraction-quality crystals were grown
by the hanging-drop vapor diffusion method at 4 'C. Crystallization solutions contained 120 mM
magnesium acetate, 50 mM sodium cacodylate pH 6.5, 1 mM spermine, and 28% w/v
polyethylene glycol (PEG) 4000; hanging drops contained 2 pL of 0.4 mM Pt-DNA in water and
106
2 pL of crystallization solution, equilibrated against 1 mL of crystallization solution. All
solutions were prepared and sterile filtered immediately before use. Crystals with dimensions of
~1.0 mm x 0.2 mm x 0.1 mm grew in clusters in approximately two weeks.
Single crystals were isolated from the clusters and transferred to a cryoprotectant solution
of 120 mM magnesium acetate, 50 mM sodium cacodylate pH 6.5, 1 mM spermine, 28% w/v
PEG 4000, and 15% v/v glycerol, mounted on loops, and flash frozen directly in liquid nitrogen.
Diffraction data were collected at 100 K (k = 0.979 A) at beamline 9-2 of the Stanford
Synchrotron Radiation Laboratory (SSRL), and processed in HKL2000. 9 Data were collected
over 360' with 10 oscillation per frame. Data collection statistics are shown in Table 4.1.
Structure determination and evaluation. Phasing of the diffraction data was achieved by
molecular replacement with the program Phaserl using the original structure (lAIO) with all
water molecules removed as a search model. After one round of restrained refinement with
Refmac5"', the Rfree value was below 24%. Platinum-nitrogen bond distances and angles were
adjusted to fit the improved electron density around the ammine ligands, and [Mg(H 2 0) 6 ]2 + sites
were inserted manually into the 2Fo-Fe and Fo-Fc electron density maps with the program Coot.' 2
Water molecules were placed automatically with Coot into areas of F0 -Fe density above 3.5y and
then adjusted manually. Subsequent rounds of refinement and model adjustment were performed
to afford the final model with an Rfree value of 19.8%. Final refinement statistics are given in
Table 4.1.
107
Table 4. 1. Data collection and refinement statistics. a
SSRL BL9-2
0.979 A
100 K
P1
Beamline
Wavelength
Collection temperature
Space group
Unit cell parameters
a
b
31.30 A
35.43 A
C
45.13 A
a
80.060
84.090
81.770
2
z
48,729
1.77 (1.83-1.77)
16734
96.0 (85.2)
2.9 (2.5)
9.2 (38.9)
8.0 (2.5)
Unit cell volume (A3 )
Resolution limit (A)
Unique reflections
Completeness (%)
Redundancy
Rmerge (%)b
I/a(1)
Resolution range (A)
R (%)c
50 - 1.77
Rfree (%)d
19.8
17.2
B-factors (A )
DNA
Water
Pt
Mg
RMSD bond lengths
RMSD bond angles
DNA atoms
Water atoms
Mg atoms
in parentheses are for the highest resolution
2
aValues
37.8
47.2
28.7
46.5
0.011 A
1.880
972
148
4
shell.
bRmerge =
II-(IAI.
= YIFo\ - |Fe\\/L|Fo|.
cR
dRfree
= R obtained for a test set of reflections (5% of diffraction data).
Nucleic acid geometric parameters were calculated with the program 3DNA. 13 Single
point energy calculations of the 1,2-cis-{Pt(NH 3)2 }2 -d(GpG) moiety were performed with the
108
Gaussian 03 suite of programs.' 4 From the experimental structures, the atomic valences were
completed with hydrogen atoms. Thus, the final model carries a molecular charge of 2+. In this
approach, the models are treated by density functional theory (DFT), with the B3LYP
functional' 5 , 16 and the 6-3 1g(d,p) basis set' 7 on all light atoms (H, C, N, 0 and P). Platinum was
represented by the Los Alamos LANL2DZ basis,1'8" 9 that includes relativistic effective core
potentials. All crystallographic images in this paper were created using Pymol.2 0 Coordinates
have been deposited into the Protein Data Bank (PDB) with accession code 3LPV.
Results and discussion
Unit cell and crystal packing. Crystals of the Pt-DNA construct were grown under different
conditions than those reported from the original investigation; however, the two structures are
nearly identical. The present conditions were used to obtain diffraction quality crystals of DNA
duplexes containing an oxaliplatin-DNA cross-link 2 ' as well as a monofunctional Pt-DNA
adduct of cis-diammine(pyridine)chloroplatinum(II)."
Despite differences in the present and
previous growth conditions, the cisplatin-modified duplex DNA crystallizes in the same space
group, P1, and with nearly identical until cell dimensions as the previous structure. Unit cell
dimensions for the original crystal structure are a = 31.27
79.810,
#l =
35.43 A, c
A3
=
A,
b = 35.46
A, c
= 47.01
A,
a =
84.75', and y = 82.79', whereas the dimensions for this crystal are a = 31.30 A, b =
45.13 A, a
and 48,729
=
80.06', fl
A3 , respectively.
=
84.09', and y = 81.77', giving unit cell volumes of 50,770
The 4% decrease in volume can be attributed to differences in
temperature; data for the first cisplatin-DNA structure were collected at 277 K, whereas the
current data were collected at 100 K. Because the unit cells are nearly equivalent, packing
interactions between molecules are also conserved between structures. Two unique DNA
109
duplexes comprise the asymmetric unit that are related by non-crystallographic symmetry and
interact via hydrogen bonding, end-to-end, and end-to-groove contacts in the crystal lattice. The
combination of the latter two interactions in the same crystal is unusual, the former being
typically observed in B-form DNA structures and the latter in crystals of A-form nucleic acids.
(a)
(b)
G6*
G6*
C19
C19
400(
C18~
(C)
C18
G23
G23
~G24G2
Figure 4.2. Structural features of cisplatin-damaged DNA. (a) Overall structure of duplex DNA
containing a cisplatin cross-link (shown in white/gray). (b) Stereo images of the platinum-bound
base pairs in molecule A with 2F0 -Fe electron density (green around the DNA/blue around the Pt
adduct) contoured at 1.5c. (c) Stereo images of a [Mg(H 2O) 6 ]2 + octahedral site bound in the
major groove of molecule A at guanine residues 23 and 24, with 2Fo-Fe electron density (shown
in blue) contoured at L.5c.
Global DNA
geometry. Notable characteristics of DNA containing the cisplatin 1,2-
{Pt(NH 3)2 }2 -d(GpG) cross-link, determined from the original crystal structure, include bending
of the double helix by 35-40' toward the major groove and local duplex unwinding of ~25'. The
roll angle between Pt-bound guanine bases is 26' (see Figs. 4.2.a and SuppL. Fig. 4.1S). DNAs
from the previous and current models align nearly identically, with a root-mean-square deviation
(rmsd) over all atoms of 0.472 A (see Fig. 4.3). Although there are two crystallographically
unique molecules in the unit cell, the DNA structures of each are also equivalent, with a rmsd
110
over all atoms of 0.18 A. Analysis of the DNA structure beyond what can be found in the
original publication6 will therefore be restricted to that which is clarified or changed after
collection of the present high-resolution data.
Figure 4.3. Stereo view of molecule A of the previously published structure of DNA modified
with a 1,2-cis-{Pt(NH 3)2}2+-d(GpG) cross-link (PDB accession code 1AIO, shown in blue)
superimposed on the current high-resolution structure (3LPV, in red). The two molecules align
almost identically, with a rmsd over all atoms of 0.472 A.
The DNA takes on A-form properties on the 5' side of the Pt-DNA adduct and is a Bform structure on the 3'-end. In the original crystal structure the resolution limit prohibited direct
visualization of deoxyribose sugar puckers, so ring conformations were indirectly determined by
measuring distances between adjacent phosphate atoms and examining base-stacking patterns.
However, in the current 1.77 A structure, sugar ring conformations are clearly differentiated as
either C3'-endo (typical of A-form DNA) or C2'-endo (for B-form DNA) in the electron density
maps (see Fig. 4.4). This unusual A/B structural feature was originally postulated to be caused
by the presence of [Co(NH 3 )6]3 + in the crystallization solution, because this complex stabilizes
111
A-form DNA.
23
This suggestion is clearly incorrect, however, because hexamminecobalt(III)
was not present in these experiments. A-form DNA occurs in several protein complexes in vivo,
including a region of the Xenopus 5S RNA gene bound by transcription factor IIIA 24 and the
DNA binding site of HIV reverse transcriptase.25 A-DNA may be preferable for protein binding
because the greater rigidity of its helix makes for a more favorable recognition site. This analysis
is consistent with a previous hypothesis that cisplatin acts on cells in part by forming DNA crosslinks that alter DNA structure to mimic protein binding sites, possibly to hijack nuclear proteins
and disrupt their function.26,27
A20
A20
C3'
C3'
G14
G14
CT
C2'
Figure 4.4. Stereo images of 2F-Fe electron density maps, shown in magenta, defining
deoxyribose sugar conformations. Top: deoxyadenosine 20, representative of A-form DNA, with
the sugar ring in C3'-endo conformation. Electron density map is contoured at 3u. Bottom:
deoxyguanosine 14, representative of B-form DNA, with the sugar ring in C2'-endo
conformation. Electron density map is contoured at 1.5-.
In the present structure, the A-like geometry is reflected by large negative slide values for
base pair steps to the 5' side of the cisplatin damage. Displacement of base pairs in this manner
causes strain between the sugar ring and phosphate group, which is relieved by flipping the 3'
112
carbon atom of the deoxyribose ring away from the phosphate into an endo conformation,
conveying A-form properties. 28 This pressure is absent to the 3' side of the helix, where the slide
values are nearly zero at each base pair step, allowing the deoxyribose moieties to adopt a Bform structure. In other structures of duplex DNA containing the 1,2-cis-{Pt(NH 3)2}2+-d(GpG)
cross-link, including an NMR solution structure2 9 and X-ray crystal structure with bound
HMGB 1 domain A, 30 the 5' platinated guanosine adopts the C3'-endo sugar pucker; however, the
remainder of the double-stranded DNA adopts B-form structure. Thus the presence of A-like
DNA in the present determination may be influenced by crystal packing interactions. Finally,
although many features of the double helix to the 5' side of the Pt cross-link resemble A-form
DNA, other geometric characteristics are more like those of B-DNA. In addition to the
previously mentioned C3'-endo sugar puckering and highly negative slide values, the A-like
segments of the duplex have average helical twist values (30.40, see Supp. Fig. 4.3S) and minor
groove widths (10.3
A)
similar to those of typical A-form (30.4' and 10.0 A, respectively)
compared to B-form DNA (35.6' and 6.2
A,
respectively).3 1 However, the average roll angle
along the same six base-pair segment (5.1 ) assumes an intermediate value between those of BDNA (1.6') and A-DNA (10.0). Similarly, the mean inclination angle (9.70) at the 5' end of
each duplex more closely resembles that of B-DNA (3.40) than A-DNA (20.10). Therefore,
although many aspects of this portion of the structure display A-form DNA features, there are
some B-DNA similarities, and nucleic acids such as the present one cannot classified as strictly
A- or B-form.
Pt adduct geometry. Unlike the overall DNA structure that is unchanged by addition of higher
resolution diffraction data, the cisplatin adduct is significantly altered in the current model. Clear
113
2
electron density is now visible for the ammine ligands of the 1,2-cis- {Pt(NH 3 )2 } +-d(GpG) cross-
link (see Fig. 4.2.b), allowing the square-planar Pt geometry to be accurately evaluated. The PtN bond lengths for all four ligands average to 2.02(4) A in the new model, compared to 1.90(1)
A in the previous structure, and 2.02(3) A in the X-ray crystal structure of the 1,2-cis32
{Pt(NH 3)2 }2+-d(GpG) dinucleotide determined at atomic resolution. The average cis N-Pt-N
angles for the current and prior models and the dinucleotide structure are 90.0', 88.0', and 90.00
respectively, with ranges of 83.90 - 99.80, 70.90 - 99.90, and 86.90 - 94.3', respectively. The
trans angles are dramatically improved in the high-resolution model, with an average N-Pt-N
value of 174.50, compared to 156.90 in the 2.6 A-resolution model and 175.10 for the isolated Pt-
d(GpG) cross-link. Thus the current model depicts a platinum adduct geometry that very nearly
2
matches that observed in the X-ray crystal structure of the 1,2-cis-{Pt(NH 3)2} +-d(GpG)
dinucleotide. A full comparison of Pt geometries is provided in Table 4.2.
Table 4.2. Comparison of platinum adduct geometries between the current high-resolution
structure, the published structure 1AIO, and the X-ray crystal structure of the platinated
dinucleotide 1,2-cis-{Pt(NH 3)2}2+-d(GpG). aAverage of two measured values from each
crystallographically unique Pt-DNA molecule. bAverage of four measured values.
Pt(GpG)T
2.05
2.04
1.98
2.02
2.02(3)
Range
2.00-2.09
2.02-2.08
1.92-2.05
1.97-209
88.1
99.9
89.0
88.8
88.0 -90.3
86.9 - 89.5
71.4
95.2
70.9-71.9
91.8-98.5
90.1
92.0
88.1 -92.2
90.9-94.3
175.0 - 177.5
155.1
154.9 - 155.2
177.0
173.3 - 179.1
171.4 - 176.5
158.8
157.1 - 160.5
175.6
174.2 - 178.1
Range
1.88- 1.89
1.90-1.92
1.86- 1.91
1.90- 1.92
3LPVa
2.03
1.99
2.00
207
2.02(4)
Range
1.98-2.07
1.88-2.10
1.98-2.02
2.06-2.08
1AIOa
1.89
1.91
1.89
1.91
1.90(1)
97.1
88.9
92.3 - 101.3
87.4 - 90.4
85.7
99.7
83.2
99.5
N7(3'G)-Pt-N2 (0 )
N1-Pt-N2 (0 )
89.0
85.1
87.7-90.4
83.7-86.5
N7(5'G)-Pt-N2 (0)
176.3
173.9
Pt-Ni (A)
Pt-N2 (A)
Pt-N7(5'G) (A)
Pt-N7(3'G) (A)
Avg. Pt-N (4)
N7(5'G)-Pt-N1 (0)
N7(5'G)-Pt-N7(3'G) (0)
0
N7(3'G)-Pt-N1 ( )
-
Consistent with the original model, the platinum atoms are displaced from the guanine
base planes. The metal atom lies out-of-plane of the 5' guanine by 1.2 and 1.1 A in the two
molecules, and of the 3' nucleobase by 1.0 A in both DNA duplexes. This feature, however, is
114
inconsistent with data from both the X-ray crystal structure of the 1,2-cis-{Pt(NH 3)2}2+-d(GpG)
dinucleotide32 and the NMR solution structure of duplex DNA containing the lesion, 29 in which
the Pt atom is located within the guanine base planes and the dihedral angle between bases is
closer to 900 (see Fig. 4.5). The shallow roll angle in the present X-ray crystal structure causes
significant angular strain on the Pt-N7 bonds. Single point energy calculations indicate that the
{Pt(NH 3)2 }2+-d(GpG) moiety in the Pt-DNA X-ray structure is 14.6 kcal/mol higher in energy
than the analogous NMR solution structure. 29 DNA end-to-end packing on the 3' side of the Pt
adduct and end-to-groove packing on the 5' side of the cross-link appear to dictate the global
curvature of the double helix and prevent complete opening of the guanine-guanine base pair roll
angle. However, the same crystal packing contacts are capable of stabilizing the altered higher
energy conformation, since a single hydrogen bonding interaction can contribute ca. 1 - 9
kcal/mol in energy.
These results underscore the importance of evaluating macromolecular
structures by multiple means, including NMR spectroscopy and biochemical methods in addition
to X-ray crystallography, because each has its limitations.
6'-dG'
3'-dG
3'-dG
(a)
3'-dG
(b)
(C)
Figure 4.5. Views of the 1,2-cis-{Pt(NH3)2} 2+-d(GpG) adduct as depicted by (a) the X-ray
crystal structure of the isolated dinucleotide,32 (b) the NMR solution structure of the Pt moiety in
8-mer duplex DNA, 29 and (c) the X-ray crystal structure determination of dodecamer DNA
presented here.
115
Magnesium sites identified. Four fully aquated magnesium(II) cations were located in the major
groove on the A-form side of the nucleic acid double helix. As in other DNA crystal structures,
interaction of the Mg(aq) 2+ complex and the DNA major groove occurs via hydrogen bonding,
the water ligands serving as hydrogen bond donors to the N7 atoms of guanine and adenine bases
and to the 06 atom of guanine (see Fig. 4.2.c). 34 '3 5 In all cases the [Mg(H 2O) 6 ]2 + cations interact
with adjacent purine bases, either the terminal d(GpG) or the d(ApG) dinucleotide sequence at
the 3' end of the unplatinated strand (see Fig. 4.6).
Significantly better electron density exists for two Mg(aq) 2+ complexes bound to the
terminal d(GpG) sequence than the other two sites. Crystallographic B-factors for the former are
38.9 and 41.0 A2 , compared to 50.5 and 51.6 A2 for the latter. The d(GpG)-bound [Mg(H 2O) 6] 2+
complexes contribute four hydrogen bonding interactions with the DNA bases, whereas the less
ordered sites participate in only three such contacts. This difference is one possible reason why
the first two sites have lower B-factors. As a result, the Mg-OH 2 bond distances for the former,
which average 2.05 ± 0.08 A, are close to the reported literature values of 2.06 A,3 and the
complexes have nearly ideal octahedral symmetry. The magnesium(II) complexes that bind
d(ApG) sequences near the platinum adduct, bases 20-21 and 44-45 as depicted in Fig. 4.7, have
Mg-OH 2 bond distances as long as 2.46 A if left unrestrained (data not shown), which is clearly
unrealistic and an artifact of insufficiently well defined electron density around those water
ligands. These complexes were therefore refined with restraints on bond distances and angles,
whereas the d(GpG) coordinated Mg(aq) 2+ sites were refined without restraints. No
[Mg(H 2 0)] 2 +-DNA interactions were observed at the B-form end of this palindromic sequence,
although they are probably present but not visible in the electron density. Average
crystallographic B-factors for the four terminal base pairs on the 5' side of the Pt cross-link,
116
where Mg(aq) 2+ coordination is resolved, are 33.8 and 36.0 A2 for the two DNA duplexes. In
contrast, these values are 45.6 and 46.0 A2 for the last four base pairs on the 3' ends of the two
molecules, indicating that the B-form sections of each nucleic acid are less ordered than the
respective A-DNA segments.
Figure 4.6. Binding of [Mg(H 2 0)6 ]2 + cations to purine dinucleotides in the cisplatin-DNA
dodecamer duplex. Hydrogen bonding interactions between water ligands and nucleobases are
depicted in cyan.
Ordered water molecules. Of the 148 ordered water molecules located in the model, 24 are
bound to magnesium in an octahedral geometry as described above. The remaining 124 solvent
molecules interact with either the platinum adduct, phosphodiester backbone, or the DNA bases.
A schematic depiction of each of these solvent-DNA interactions is given in Fig. 4.7, and a full
list of water contacts appears in SuppL. Table 4.1S. One water molecule participates in bridging
hydrogen-bonding interactions with each of the platinum ammine ligands. Molecule A has four
solvent molecules bound to the Pt-DNA adduct, whereas molecule B has only two ammine-
117
bound waters. Unlike another high-resolution structure of Pt-modified DNA, no direct
interaction between ordered waters and the platinum atom is observed.3 6 3 7
Hydrogen bonding interactions between water molecules and nucleobases occur
primarily at guanine N7, 06, or N3 atoms, adenine N7, N6, or N3, thymine 02 or 04, and
cytosine N4 or 02 positions, consistent with previous observations of Watson-Crick base pair
hydration patterns.3 8 ,3 Water contacts with oxygen atoms of the deoxyribose rings (03', 04', or
05') are also observed, although less regularly. There are more resolved water-nucleobase
interactions at the central base pairs in the duplexes; terminal base pairs do not show as much
detail in the hydration sphere, presumably due to increased thermal motion.
5'
3 '-HH
1
CO
~
24,
.
23
MOM 108--HOH 35
MOM..
HOH27
2
CG
HOH 131--
3
T
HOH129
MOM 125--._
HOH 79
HH7OH
HH7-,HO
HO
HOH
O--MOM
H.-H" 80 96
HOH 5
6
HOH
HOH 113
136
HO
18
917
91OH
5
10
MOM 30
12
HHIl
3'
G
CG
14
T~
46
29
HOH 121
OH 14-
['MHOH
HOH 92
H HOH
(29
32
45
T
5
[M (HOH)
HOM 11
44
2OH
31
HOH 22MOM 72MOM 84 ;P3
-3-
MOM 46
MOM 49 O
10
MOM19
MOM 38
OM 13
2
MO
OM 112
3
35
36
13
3'
5'
MOM 88
HOH 86
42
83
MOM 92
HOH 3
7
O10
41
MOM 122
MOM 119
HOH 135
MOM 128
CG
40
TA
39
CG
CG
8
HO3
26
POH
H
HOH 26
HOH 89-OM 87
HOH
asM C
G0 4
MO 1
OH 71MOM 1168
j
-MOH 67
HOH 102
HOH 36-HOH 115
MOH 120360-OH11
HH1
TA
1
47
28
97
12 1
HOHH14--
39
HOH 971
MOM 18MOM 100--1
HO 2 1
127
HOH 95
CG
O M 10
HMH 7-
H
HOH 53
MOM 76
,.-HOH134z
109
OM 93
M9
OM 101
is
CHOH
WI
MOM 932
97
27
HOH 10
-HOH
HO
H20
1
P
41-._.
,'MOH
HOH 17HOM 28
26
19
7
77
H2
- HO H 44
20M
H1MOM 131M
OH
OM110
20
9
HOH 78
MOM
O 51
MOH 66
HOH
5"4-69.
O M3
HOH 52
3'
2518[g(HM 48
HOH
OH 82-.._
O 81
HOH40
HOH 103
450
H1H
OH
118
5'
994
H
42CG
HOH 43
OM 3
[Mg (HOH)
HOH 16
O
38
1
MM3
-MOM
37
123
M068
5'
Figure 4.7. Schematic depicting hydrogen-bonding interactions between solvent molecules and
Pt-DNA. For clarity, many contacts between water molecules are omitted.
118
The phosphodiester backbone of each DNA double helix is extensively hydrated, with
water molecules participating in hydrogen bonding with the phosphate oxygen atoms. Mono- and
bidentate interactions, either with two oxygen atoms from a single phosphate or from adjacent
nucleotides, occur. Most bridging interactions between two phosphates are found in the central
four base pairs, where the DNA backbone structure is distorted by the cisplatin cross-link. Some
of the atoms modeled here as oxygen from water may actually be sodium ions, given that 50 mM
sodium cacodylate was present in the crystallization buffer and that DNA exists as a polyanion
and must be neutralized by cations. However, these two species cannot be differentiated by Xray crystallography since Na+ and 02- each contain ten electrons; therefore, all non-magnesium
solvent molecules were modeled as water.
Conclusions
The structure of a dodecamer DNA duplex containing a centrally located 1,2-cis{Pt(NH 3)2 }2+-d(GpG)
crystallography to 1.77
intrastrand
A
cross-link
of
cisplatin
resolution, a greater than 0.8
A
was
determined
by
X-ray
improvement over the previous
structure solved in 1996. This improvement in resolution advances understanding of the structure
of cisplatin-modified duplex DNA in four primary areas: (i) deoxyribose sugar puckers are
identified directly from the electron density and compare the resulting A/B DNA hybrid to other
Pt-DNA structures and to biologically relevant non-standard DNA geometries; (ii) the structure
of the platinum adduct is more accurately determined, and now closely matches idealized squareplanar geometry, eliminating the possibility that the DNA duplex distorts the structure of the
primary coordination sphere; (iii) the characteristic features of the {Pt(NH 3)2 }2+-d(GpG) moiety
119
are sharpened, specifically with regard to the guanine plane orientation; and (iv) four Mg(aq)2 +
complexes and dozens of additional interacting solvent molecules are identified.
Knowledge of the structural changes to DNA that occur upon binding of platinum
antitumor agents is critical in deciphering the mechanism of these compounds. X-ray
crystallography is a powerful technique for answering some of these questions, and should
continue to be relied upon in the future as other platinum complexes are synthesized and studied.
From these data one can better understand how Pt-DNA adducts might be repaired and inhibit
cellular transcription. Understanding these two functions is key to the design of improved
platinum candidates for cancer chemotherapy.
120
References
(1) Kelland, L. Nat. Rev. Cancer 2007, 7, 573-584.
(2)
Rosenberg, B.; Van Camp, L.; Krigas, T. Nature 1965, 205, 698-699.
(3) Rosenberg, B.; Van Camp, L.; Trosko, J. E.; Mansour, V. H. Nature 1969, 222, 385-386.
(4)
Wang, D.; Lippard, S. J. Nat. Rev. Drug Discovery 2005, 4, 307-320.
(5) Takahara, P. M.; Rosenzweig, A. C.; Frederick, C. A.; Lippard, S. J. Nature 1995, 377,
649-652.
(6)
Takahara, P. M.; Frederick, C. A.; Lippard, S. J. J. Am. Chem. Soc. 1996, 118, 1230912321.
(7)
Todd, R. C.; Lippard, S. J. J Inorg. Biochem. 2010, 104, in press.
(8) Dhara, S. C. Indian J Chem. 1970, 8, 193-194.
(9)
Otwinowski, Z.; Minor, W. Methods Enzymol. 1997, 276, 307-326.
(10) McCoy, A. J.; Grosse-Kunstleve, R. W.; Adams, P. D.; Winn, M. D.; Storoni, L. C.; Read,
R. J. J. Appl. Cryst. 2007, 40, 658-674.
(11) Murshudov, G. N.; Vagin, A. A.; Dodson, E. J. Acta Crystallogr.1997, D53, 240-255.
(12) Emsley, P.; Cowtan, K. Acta Crystallogr.2004, D60, 2126-2132.
(13) Lu, X.-J.; Olson, W. K. Nucleic Acids Res. 2003, 31, 5108-5121.
(14) Frisch, M. J.; G. W. Trucks, H. B. S., G. E. Scuseria,; M. A. Robb, J. R. C., J. A.
Montgomery, Jr., T. Vreven,; K. N. Kudin, J. C. B., J. M. Millam, S. S. Iyengar, J. Tomasi,;
V. Barone, B. M., M. Cossi, G. Scalmani, N. Rega,; G. A. Petersson, H. N., M. Hada, M.
Ehara, K. Toyota,; R. Fukuda, J. H., M. Ishida, T. Nakajima, Y. Honda, 0. Kitao,; H.
Nakai, M. K., X. Li, J. E. Knox, H. P. Hratchian, J. B. Cross,; V. Bakken, C. A., J.
Jaramillo, R. Gomperts, R. E. Stratmann,; 0. Yazyev, A. J. A., R. Cammi, C. Pomelli, J. W.
Ochterski,; P. Y. Ayala, K. M., G. A. Voth, P. Salvador, J. J. Dannenberg,; V. G.
Zakrzewski, S. D., A. D. Daniels, M. C. Strain,; 0. Farkas, D. K. M., A. D. Rabuck, K.
Raghavachari,; J. B. Foresman, J. V. 0., Q. Cui, A. G. Baboul, S. Clifford,; J. Cioslowski,
B. B. S., G. Liu, A. Liashenko, P. Piskorz,; I. Komaromi, R. L. M., D. J. Fox, T. Keith, M.
A. Al-Laham,; C. Y. Peng, A. N., M. Challacombe, P. M. W. Gill,; B. Johnson, W. C., M.
W. Wong, C. Gonzalez, and J. A. Pople, ; Gaussian, Inc.: Wallingford CT 2004.
(15) Lee, C.; Yang, W.; Parr, R. G. Phys. Rev. B 1988, 37, 785-789.
(16) Becke, A. D. J. Chem. Phys. 1993, 98, 5648-5652.
(17) Hehre, W. J.; Ditchfield, R.; Pople, J. A. J Chem. Phys. 1972, 56, 2257-2261.
121
(18) Hay, P. J.; Wadt, W. R. J. Chem. Phys. 1985, 82, 299-3 10.
(19) Hay, P. J.; Wadt, W. R. J. Chem. Phys. 1985, 82, 270-283.
(20) DeLano, W. L. DeLano Scientific, PaloAlto, CA, USA 2002.
(21) Spingler, B.; Whittington, D. A.; Lippard, S. J. Inorg. Chem. 2001, 40, 5596-5602.
(22) Lovejoy, K. S.; Todd, R. C.; Zhang, S.; McCormick, M. S.; D'Aquino, J. A.; Reardon, J. T.;
Sancar, A.; Giacomini, K. M.; Lippard, S. J. Proc. Natl. Acad Sci. US.A. 2008, 105, 89028907.
(23) Gao, Y.-G.; Robinson, H.; van Boom, J. H.; Wang, A. H.-J. Biophys. J. 1995, 69, 559-568.
(24) Fairall, L.; Martin, S.; Rhodes, D. EMBO J. 1989, 8, 1809-1817.
(25) Jacobo-Molina, A.; Ding, J.; Nanni, R.; Clark Jr., A. D.; Lu, X.; Tantillo, C.; Williams, R.;
Kamer, G.; Ferris, A.; Clark, P.; Hizi, A.; Hughes, S.; Arnold, E. Proc. Natl. Acad Sci.
U.S.A. 1993, 90, 6320-6324.
(26) Vichi, P.; Coin, F.; Renaud, J.-P.; Vermeulen, W.; Hoeijmakers, J. H. J.; Moras, D.; Egly,
J.-M. The EMBO Journal1997, 16, 7444-7456.
(27) Cullinane, C.; Mazur, S. J.; Essigmann, J. M.; Phillips, D. R.; Bohr, V. A. Biochemistry
1999, 38, 6204-6212.
(28) Dickerson, R. E.; Ng, H.-L. Proc.Natl. Acad Sci. US.A. 2001, 98, 6986-6988.
(29) Marzilli, L. G.; Saad, J. S.; Kuklenyik, Z.; Keating, K. A.; Xu, Y. J. Am. Chem. Soc. 2001,
123, 2764-2770.
(30) Ohndorf, U.-M.; Rould, M.; He,
Q.; Pabo, C.; Lippard, S. J. Nature 1999,
399, 708-712.
(31) Ng, H.-L.; Kopka, M. L.; Dickerson, R. E. Proc. Natl. Acad Sci. US.A. 2000, 97, 20352039.
(32) Sherman, S. E.; Gibson, D.; Wang, A. H.-J.; Lippard, S. J. J. Am. Chem. Soc. 1988, 110,
7368-7381.
(33) Bandyopadhyay, D.; Bhattacharyya, D. Biopolymers 2006, 83, 313-325.
(34) Soler-L6pez, M.; Malinina, L.; Subirana, J. A. J Biol. Chem. 2000, 275, 23034-23044.
(35) Subirana, J. A.; Soler-Lopez, M. Ann. Rev. Biophys. Biomol. Struct. 2003, 32, 27-45.
(36) Coste, F.; Malinge, J.-M.; Serre, L.; Shepard, W.; Roth, M.; Leng, M.; Zelwer, C. Nucleic
Acids Res. 1999, 27, 1837-1846.
(37) Coste, F.; Shepard, W.; Zelwer, C. Acta Crystallogr.2001, D58, 431-440.
122
(38) Schneider, B.; Cohen, D. M.; Schleifer, L.; Srinivasan, A. R.; Olson, W. K.; Berman, H. M.
Biophys. J 1993, 65, 2291-2303.
(39) Berman, H. M. Biopolymers 1997, 44, 23-44.
123
RISE
SHIFT
-+-
2.00
NEW mol A
3.90
NEW mol B
OLD mol A
-+-OLD mot B
---
1.50
1.00
-+-
RISE old mol B
3.50
5
3.30
0.00
3.10
-0.50
2.90
-1.00
2.70
2.50
-1.50
2
8
6
4
10
0
12
6
Base pair step
SLIDE
TILT
8.00 I
-4- SLIDE mol A
-U- SLIDE mol B
SLIDE old mol A
-A-
4
2
Base pair step
0.50
-0.50
RISE mol A
RISE mol B
RISE old mot A
3.70
0.50
0.00
----
6.00
4.00
SLIDE old mol B
8
10
12
-+-TILT mol A
-U-TILT mol B
TILT old mol A
+(- TILT old mol B
2.00
0 -1.00
0.00
-1 50
-2.00
-2.00
-4.00
-
-2.50
0
-
2
-
-
-6.00
-
4
8
6
10
12
0
2
4
6
TWIST
ROLL
35.00
10
-+--TWIST
-U-TWIST
TWIST
-+-TWIST
45.00
30.00
+- ROLLmol A
ROLL mol B
A
ROLLold mot
--X- ROLLold mol B
25.00
8
12
Base pair step
Base pair step
mol A
mol B
old mol A
old mol B
40.00
-U-
6 20.00
35.00
15 00
a
j30.00
10.00
25.00
5.00
0.00
20.00
0
2
4
6
8
10
12
2
Base pair step
4
6
8
10
12
Base pair step
Figure 4.1S. Graphical depiction of base pair step parameters for molecules A and B of the
current high-resolution structure (shown in blue and red, respectively), and the published 2.6 A
structure (yellow and green, respectively).
124
Table 4. IS. Hydrogen-bonding interactions between water molecules. Waters 54-65 and 137148, shown in boldface italics, are ligands in {Mg(H 2O)6 } 2+ complexes.
Water
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
Residue
G30
G31
G30
G31
HOH73
G31
G30
T29
T8
16G
HOH80
HOH76
G7
G6
HOH77
G6
G6
T32
G40
T5
HOH37
C28
T29
HOH69
G45
HOH138
HOH139
HOH135
C33
HOH72
HOH31
A20
HOH37
HOH103
G30
HOH15
C4
HOH14
HOH50
HOH51
G23
T8
HOH77
T1O
HOH96
HOH132
A15
CPT50
CPT50
HOH121
Atom
01P
02P
05'
02P
HOH
06
06
04
04
06
HOH
HOH
02P
02P
HOH
01P
01P
04
06
02
HOH
02
04'
HOH
02P
HOH
HOH
HOH
N4
HOH
HOH
N3
HOH
HOH
01P
HOH
02
HOH
HOH
HOH
N2
02P
HOH
02P
HOH
HOH
N6
NI
N2
HOH
125
Distance (A)
2.6
2.9
3.2
2.6
2.9
3.1
2.9
2.8
2.4
2.7
3.1
3.1
2.7
3.1
2.5
3.0
2.8
2.6
3.0
2.6
2.8
2.7
3.2
3.0
2.6
3.2
2.6
3.1
3.0
2.7
2.8
2.8
2.9
2.9
3.1
2.8
2.5
2.8
2.7
2.9
3.0
2.7
3.0
2.5
2.6
2.9
3.0
3.0
3.2
2.4
Water
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
Residue
C9
HOH41
T32
HOH71
C43
A44
G24
HOH46
HOH56
A17
HOHIO
A43
A44
HOH138
C2
T8
C28
T1O
C1l
HOH33
T34
HOH12
G45
G16
HOH30
A39
C2
G47
HOH115
HOH9
HOH13
A15
A17
C26
T8
HOH21
C1l
HOH100
G24
G21
HOH66
A20
C19
HOH148
C33
HOH24
HOH54
T29
T5
HOH118
T34
G21
HOH15
A22
HOH15
Atom
N4
HOH
02P
HOH
OlP
02P
02P
HOH
HOH
N7
HOH
02P
02P
HOH
N4
01P
N4
02
04'
HOH
04
HOH
N3
N3
HOH
N6
OlP
N2
HOH
HOH
HOH
02P
N3
N4
02P
HOH
N4
HOH
02P
02P
HOH
02P
02P
HOH
OlP
HOH
HOH
02P
04
HOH
02P
N2
HOH
N3
HOH
126
Distance (A)
2.9
2.6
2.6
2.4
2.7
2.8
2.6
2.9
3.0
2.7
3.0
3.2
3.0
3.1
2.8
2.7
2.9
2.7
3.2
2.7
2.8
2.8
3.0
2.7
2.7
2.7
3.1
3.1
2.9
2.8
2.9
2.4
2.8
3.0
2.8
2.6
2.8
3.0
2.6
3.1
2.6
2.9
3.0
3.1
2.8
2.9
2.5
3.0
3.0
2.5
2.7
3.0
2.7
2.6
2.9
Water
52
53
54
55
56
57
58
59
60
61
63
64
65
66
67
68
69
70
71
72
73
74
75
76
Residue
A22
G23
HOH54
HOH57
CPT49
HOH74
HOHIO
HOH46
HOH52
G23
G24
G24
HOH24
G23
HOH52
HOH66
HOH85
HOH66
G48
HOH67
G47
G48
G47
HOH99
HOH92
A22
HOH44
HOH57
HOH59
HOH60
G48
HOH99
G37
T5
HOH1O
HOH70
HOH69
G31
HOH22
T32
HOH12
HOH106
CPT50
HOH2
CPT49
T5
HOH75
HOH53
G6
CPT49
HOH74
HOH76
CPT49
HOH4
Atom
02P
02P
HOH
HOH
N2
HOH
HOH
HOH
HOH
06
06
N7
HOH
N7
HOH
HOH
HOH
HOH
N7
HOH
N7
06
06
HOH
HOH
N7
HOH
HOH
HOH
HOH
02P
HOH
06
o1P
HOH
HOH
HOH
02P
HOH
02P
HOH
HOH
N2
HOH
NI
02P
HOH
HOH
02P
NI
HOH
HOH
N2
HOH
127
Distance (A)
2.8
3.0
3.2
2.9
3.0
3.1
3.0
2.5
3.2
2.5
2.7
2.7
3.0
2.8
2.9
3.2
2.8
3.0
2.6
2.7
2.6
2.8
2.6
2.5
3.1
2.9
2.6
3.2
3.0
2.7
2.7
2.5
3.2
2.8
3.0
2.9
2.9
3.0
2.4
2.8
2.7
2.9
3.1
2.9
2.7
2.7
2.7
3.1
2.7
3.2
2.7
3.2
2.7
3.2
Water
77
78
79
80
81
82
83
84
85
86
87
88
89
90
91
92
93
94
95
96
97
98
99
100
101
Residue
HOH80
HOH77
HOH5
HOH17
HOH76
G7
G6
T5
HOHIO
C4
HOH4
HOH76
C26
C26
HOH90
G48
T32
HOH85
HOH106
C33
HOH84
HOH58
HOH88
HOH89
A44
HOH86
A44
HOH87
HOH82
C43
HOH98
A46
HOH65
HOH133
A17
A17
G24
C28
C9
HOH18
T1O
C9
HOH102
C43
HOH91
HOH142
C9
HOH67
HOH64
T1O
HOH42
A17
HOH25
HOH53
HOH78
Atom
HOH
HOH
HOH
HOH
HOH
06
06
04
HOH
02P
HOH
HOH
02P
OlP
HOH
03'
OlP
HOH
HOH
OlP
HOH
HOH
HOH
HOH
01P
HOH
N3
HOH
HOH
02P
HOH
N7
HOH
HOH
05'
02P
03'
o1P
02P
HOH
o1P
03'
HOH
N4
HOH
HOH
o1P
HOH
HOH
02P
HOH
N6
HOH
HOH
HOH
128
Distance (A)
3.0
2.4
2.5
3.1
2.4
3.0
2.8
2.8
3.2
2.9
3.1
3.0
2.9
3.1
3.2
2.6
2.7
2.7
2.7
2.9
2.7
2.8
3.2
2.7
3.0
3.2
2.8
2.7
3.2
2.9
3.1
3.1
3.1
2.7
3.1
3.2
3.2
2.8
2.7
2.6
2.6
3.1
2.7
2.9
3.1
2.6
3.0
2.5
2.5
2.8
3.0
3.0
3.0
3.0
3.2
Water
102
103
104
105
106
107
108
109
110
111
112
113
114
115
116
117
118
119
120
121
122
123
124
125
126
127
128
129
130
131
132
133
Residue
G48
HOH97
G21
HOH13
G21
HOH144
T27
C28
C33
HOH72
HOH84
C42
C2
C18
C19
HOH145
C12
C35
HOH126
T27
HOH137
T5
HOH6
HOH36
G30
HOH121
G40
C4
HOH48
A41
HOH128
A20
CPT50
T29
HOH20
HOH116
A41
G38
G14
A15
C4
T34
HOH112
HOH136
C28
A41
HOH119
T3
T34
C35
T3
C9
HOH18
G45
HOH92
Atom
02P
HOH
04'
HOH
02P
HOH
02P
02P
02P
HOH
HOH
OlP
02P
02P
N4
HOH
N4
N4
HOH
04
HOH
OlP
HOH
HOH
02P
HOH
N2
N4
HOH
N6
HOH
OlP
NI
02P
HOH
HOH
N3
OP
N3
04'
OlP
02P
HOH
HOH
01P
N7
HOH
02
02
04'
o1P
OlP
HOH
02P
HOH
129
Distance (A)
2.7
2.7
3.2
2.9
2.5
2.7
3.1
2.9
2.8
2.9
2.7
3.0
2.9
2.6
3.0
2.6
2.8
2.9
2.8
2.4
3.0
2.7
2.3
2.9
2.6
2.6
3.0
3.0
2.5
3.1
2.8
2.7
2.7
2.8
2.4
2.6
2.7
3.2
3.1
3.2
3.0
2.7
2.8
2.8
3.2
2.8
2.8
2.5
2.5
3.0
3.1
2.7
2.9
2.9
2.7
Water
134
135
136
137
138
139
142
144
145
147
148
Residue
C18
A44
HOHI1
HOH127
G45
HOH113
HOHI
HOH26
G45
HOHI
A44
HOH98
G21
HOH104
A20
HOHIO
G21
HOH45
Atom
OlP
02P
HOH
HOH
06
HOH
HOH
HOH
N7
HOH
N7
HOH
N7
HOH
N7
HOH
06
HOH
130
Distance (A)
3.2
3.1
3.1
2.8
2.8
3.0
3.2
3.1
2.8
2.6
2.5
2.6
2.8
2.7
2.7
2.6
2.7
3.1
Chapter 5. Structural Investigations of a Site-Specifically Platinated Nucleosome Core
Particle Containing the Cisplatin 1,3-{Pt(NH 3 )2 } 2 -d(GpTpG) Cross-link
131
Introduction
DNA adducts of cisplatin have been thoroughly characterized by a variety of biochemical
and biophysical methods, including X-ray crystallography and NMR spectroscopy, as previously
reviewed.1- 3 Structures of the 1,2-d(GpG) and 1,3-d(GpTpG) intrastrand cross-links formed
between cisplatin and duplex DNA have been solved by at least one of the above techniques.- 7
Interstrand cross-links8 and monofunctional Pt-DNA adducts9 have also been characterized by
these methods. The results have provided valuable structural information about DNA adducts of
platinum antitumor agents and yielded insight into downstream cellular processes such as DNA
nucleotide excision repair and inhibition of transcription by RNA polymerase 11.2,3 Many of the
platinum compounds being synthesized as potential anticancer drugs are direct analogs of
cisplatin. They bind DNA in a similar manner and cause analogous structural changes.
One shortcoming of all such structural work is its failure to reproduce a key component
of the eukaryotic cellular environment of nuclear DNA, namely, the nucleosome. DNA is
packaged as chromatin, the building block of which is the nucleosome core particle (NCP).
These protein-DNA complexes include ~146 base pairs of DNA wrapped in one and threequarter turns as a left-handed superhelix around a core of eight histones comprising two copies
each of H2A, H2B, H3, and H4.10 Chromatin proteins are responsible for two seemingly
contradictory functions; (i) to provide a framework for condensing 2-3 meters of human DNA
into a compact structure that fits inside the nucleus of the cell, and (ii) to regulate DNA access by
proteins involved in replication, transcription, recombination, and repair." The location of
nucleosomes in vivo is primarily regulated by the intrinsic DNA sequence. It is estimated that
over 50% of nucleosome positions are directly determined by such sequence.' 2
132
Nucleosome core particles have been well characterized by X-ray crystallography
3
-
there are currently over 30 nucleosome structures in the Protein Data Bank. The most detailed is
4
a 1.9 A resolution structure of a nucleosome containing histones from Xenopus laevis.1 Histone-
DNA contacts are observed each time the DNA minor groove faces the histone core, stabilizing
the distorted structure of the nucleic acid around the octamer. In total -120 histone-DNA
contacts are identified, nearly 50% of which involve hydrogen bonding interactions between the
phosphate backbone of DNA and the protein main chain. No interactions occur between the
histone core proteins and the DNA nucleobases. Furthermore, an additional -120 solventmediated interactions exist, whereby water molecules form bridging hydrogen-bonding contacts
between protein and DNA. These observations suggest that DNA positioning sequences organize
chromatin based on geometric properties of the duplex dictated by nucleotide sequence and not
by direct control with the bases. In this manner the histone core can readily package DNA of any
sequence into nucleosomes. Another conserved feature of nucleosome structure is the presence
of arginine side chains that form contacts between the histone octamer and DNA at 12 of the 14
minor groove locations. N-terminal histone tails extend around and through the DNA double
helix, creating additional protein-DNA interactions, but are disordered in most X-ray crystal
16
structures. These features are generally conserved in nucleosomes from X laevis,is S. cervisae,
D. melanogaster,7 and human sources,' 8 due in large part to the extensive sequence homology
between histone proteins.19
Although nucleosomes are inherently stable protein-DNA complexes, mechanisms exist
that permit access by cellular proteins to the underlying DNA. The histone octamer is
translocated along DNA strands by either ATP-independent or -dependent pathways. In the
former process, nucleosome sliding occurs in a temperature-dependent manner that reflects the
133
stability of histone-DNA interactions for a given nucleosome. 19,20 In vivo nucleosome
reorganization is directed primarily by ATP-dependent chromatin remodeling complexes 2 ' and
histone chaperones. 22 These processes are reviewed elsewhere, but the mechanism of chromatin
remodeling may involve the previously mentioned arginine residues that protrude into the minor
groove.
Proper nucleosomal positioning and mobility are critical to the fidelity of eukaryotic
transcription."
Initial transcription factor binding occurs at DNA promoter sites that are
characteristically nucleosome-free, which allows the proteins to recognize and bind the exposed
DNA sequence. As the RNA polymerase elongation complex transcribes along the DNA
template, upstream nucleosomes are continually shifted and unwrapped by chromatin remodeling
complexes such as FACT.
Despite the significant investment in studying interactions between cisplatin and DNA,
very few reports exist in the literature discussing effects of platinum antitumor drug binding to
chromatin or nucleosomes. Early studies revealed that cis- and trans-diamminedichloroplatinum(II) bind histone-bound and linker DNA with nearly equal affinity,25,26 with a minor
preference for linker DNA at low platinum concentrations.27 2 8 Cisplatin forms primarily DNA
intrastrand cross-links with nucleosomes, whereas the trans isomer forms primarily histonehistone and histone-DNA cross-links. 2 5 NMR spectroscopic evidence suggests that the crosslinks formed with free and nucleosomal DNA by cisplatin are equivalent, 29 without significantly
altering the helical twist required for DNA packaging in the nucleosome.2 7 An isolated report
presents conflicting data that cisplatin treatment of nucleosomes causes dissociation of DNA
from the histone core, whereas trans-diamminedichloroplatinum(II)
causes no effect on
nucleosome stability. 30 However, these experiments utilized closed circular DNA and high
platinum concentrations, so the physiological relevance of this result is questionable.
134
More recent studies of platinum-nucleosome interactions have focused on structural
effects of cisplatin binding to DNA wrapped around histone complexes. Data from chemical
footprinting experiments demonstrate that DNA containing a site-specific cisplatin 1,2-d(GpG)
or 1,3-d(GpTpG) intrastrand cross-link enforces a characteristic rotational orientation of the
DNA strand on the nucleosome, such that the Pt adduct faces inward toward the histone core.
1-33
Such platinum-DNA cross-links are repaired less efficiently from nucleosomal compared to free
DNA. 34 These structural insights suggest a mechanism by which platinum damage can be
shielded from repair by the nucleosome surface. Other results indicate that platinum damage
does not significantly affect the translational positioning of nucleosomes. In these experiments
nucleosome core particles were treated with cisplatin or oxaliplatin, and both electrophoretic
mobility shift assays and X-ray crystallography, involving nucleosome crystals treated with
either drug, indicated no changes in DNA translational position.3 s,3 6 The difference between the
two sets of experiments is that the former results describe formation of nucleosomes from
platinated DNA, whereas the latter involve platinum treatment of pre-assembled nucleosomes.
Together, these studies suggest a mechanism whereby platinum binds nucleosomal DNA in
positions where the adduct is most readily accommodated by the nucleosome structure,
reinforcing the native positioning preference instead of modifying it. Enhanced cisplatin binding
37
at bent DNA sites caused by protein binding has been previously encountered.
Despite the recent work, significant questions remain. How do platinum intrastrand crosslinks determine nucleosome rotational phasing? To what extent is the global architecture of
NCPs affected by cisplatin-DNA damage? In order to explore these issues, the X-ray crystal
2
structure of a nucleosome core particle containing a single {Pt(NH 3)2}
modification was
determined. This report describes the preparation, purification, crystallization, and 3.2 A X-ray
135
structure determination of nucleosomes prepared from recombinant histones from X laevis and a
synthetic 146-bp DNA containing a site-specific l,3-cis-{Pt(NH 3)2}2 -d(GpTpG) intrastrand
cross-link. This adduct is commonly thought to be the major adduct of carboplatin, 38 and is more
efficiently repaired than the corresponding 1,2-d(GpG) cross-link. Given the eukaryotic cellular
environment and typical platination levels of DNA in cancer cells, the mononucleosome model
containing a single Pt-DNA cross-link provides the most physiologically relevant information
about of cisplatin-DNA modification to date.
Experimental
Materials. Phosphoramidites, columns, and other reagents for solid-phase oligonucleotide
synthesis were purchased from Glen Research. Potassium tetrachloroplatinate(II) used to prepare
cisplatin 39 was a gift from Engelhard Corporation (now BASF). Enzymes were purchased from
New England Biolabs. y-32P-ATP (6000 Ci/mmol) was procured from Perkin Elmer. All other
reagents were purchased from commercial suppliers and used without further purification. UVVis spectroscopy was performed with a Hewlett-Packard 8453 instrument, and liquid
chromatography with an Agilent 1200 series HPLC equipped with a temperature-controlled
autosampler and automated fraction collector. Gel filtration chromatography was performed with
an Akta FPLC instrument at 4 'C. MALDI mass spectrometry was conducted on a Bruker
Omniflex instrument. All dialyses were performed using Spectra/Por dialysis membranes of an
appropriate molecular weight cut-off and were pre-treated with hot 50 mM aqueous EDTA,
followed by several washes with water and dialysis buffer, prior to use. Atomic absorption
spectra, used to quantitate platinum concentrations, were recorded with a Perkin Elmer AAnalyst
300 system. Radioactive gels were visualized using a Storm 840 phosphorimager, and sample
136
radioactivity was quantitated with a Beckman LS 6500 scintillation counter. Syntheses of duplex
t2-Pt (Fig. 5.1) and the corresponding nucleosome core particle were conducted in collaboration
with the technical assistance of Paresh Agarwal.
Oligonucleotide synthesis. Oligonucleotides v, x, yl, y2, and z, shown in Fig. 5.1, were
synthesized on a 1.0 pmol scale, deprotected overnight at 60 'C, dried in a vacuum centrifuge,
and purified by 4% denaturing gel electrophoresis. Following extraction from the gel, samples
were ethanol precipitated, desalted with SepPak C18 solid-phase extraction cartridges, and
40
quantitated by UV-Vis spectroscopy using calculated extinction coefficients. Oligonucleotides
w1 and w2 were synthesized with dimethoxytrityl (DMT) groups on and purified by semipreparative HPLC on an Agilent SB-300, 9.4 x 250 mm column using method RCT.005S,
described in Appendix C. After lyophilization of combined fractions, DMT groups were
removed in 80% acetic acid for 30 min at room temperature, and the oligonucleotides were
precipitated with isopropanol, desalted with Sep-Pak C18 cartridges, and quantitated by UV-Vis
spectroscopy. From these strands, wl-Pt and w2-Pt were prepared by incubation of the starting
materials with 1.2 equiv of cisplatin in 10 mM HEPES pH 6.8 for 14 h at 37 'C according to
published procedures. 4 1 The former product was purified by ion-exchange HPLC on a Dionex
DNA-Pac PA-100, 9.4 x 250 mm column using method RCT.006S, and the latter platinated
strand by method RCT.007A (both detailed in Appendix C). Pure fractions were dialyzed
against water overnight, then lyophilized, reconstituted in water, and quantitated by UV-Vis and
AA spectroscopy. Yields for platination reactions ranged from 37 to 44%. Pt/DNA ratios for wlPt and w2-Pt ranged from 1.03 - 1.15, indicating 1 Pt atom per DNA strand. HPLC
chromatograms showing purification of each platinated strand are presented in Figs. 5.2 and 5.3.
137
(a)
v (63mer):
5 'ATCAATATCCACCTGCAGATTCTACCAAAAGTGTATTTGGAAACTGCTCCATCAAAAGGCATG
wl (14mer):
5'TTCACCGTGATTCC
wl-Pt (Pt-14mer):
5'TTCACCGTGATTCC
w2 (14mer):
5'TTCACCGGAATTCC
w2-Pt (Pt-14mer):
5 'TTCACCGGAATTCC
x (69mer):
5'CCTCAACATCGGAAAACTACCTCGTCAAAGGTTTATGTGAAAACCATCTTAGACGTCCACCTA
TAACTA
yl (86mer):
5 'ATGTTGAGGGGAATCACGGTGAACATGCCTTTTGATGGAGCAGTTTCCAAATACACTTTTGGT
AGAATCTGCAGGTGGATATTGAT
y2 (86mer):
5 'ATGTTGAGGGGAATTCCGGTGAACATGCCTTTTGATGGAGCAGTTTCCAAATACACTTTTGGT
AGAATCTGCAGGTGGATATTGAT
z (60mer):
5'TAGTTATAGGTGGACGTCTAAGATGGTTTTCACATAAACCTTTGACGAGGTAGTTTTCCG
(b)
v
w(Px
t(Pt
ligation
Figure5.1. (a) Oligonucleotides synthesized towards ligation of 146 bp DNA containing single
cisplatin cross-links. Bases shown in red depict platination sites. (b) The 146 bp ligation scheme.
90
80
70
60
50
40
30
20
10
00
5
10
15
20
25
30
35
40
45
Time (min)
Figure5.2. HPLC purification of wl-Pt (retention time ~ 23 min) from side products.
138
2500
2000
1500
1000
500
0
5
10
15
20
25
30
35
40
45
Time (min)
Figure 5.3. HPLC purification of w2-Pt (retention time ~ 12 min) from side products.
Characterization of wl-Pt and w2-Pt: MALDI-TOF mass spectrometry. An aliquot of each
oligonucleotide (0.5 pL) was mixed with 20 ptL of 10 mg/mL 2',4',6'-trihydroxyacetophenone
monohydrate in 25 mM ammonium citrate, 50:50 MeCN:H 20. The samples were analyzed on a
Bruker Omniflex MALDI-TOF mass spectrometer in negative ion mode. Masses for wl-Pt and
w2-Pt; calculated: 4416.0 Da and 4428.0 Da, respectively. Found: 4417.1 Da and 4424.7 Da,
respectively.
Characterization of wl-Pt and w2-Pt: Nuclease Sl/CIP digestion. In separate Eppendorf
tubes, 2 nmol each of wl-Pt and w2-Pt were incubated with 5 pL Sl nuclease in 100 ptL of
reaction buffer (50 mM sodium acetate pH 4.5, 280 mM NaCl, 4.5 mM ZnSO 4) at 37 'C
overnight. To these samples were added 5 pL 1.5 M Tris-HCl pH 8.8 and 1 pL calf intestinal
phosphatase; the solutions were incubated for another 4 h at 37 0 C. After addition of 6 ptL 0.1 N
HCl and centrifugation, the samples were analyzed by HPLC (see RCT.008A, Appendix C).
Calculated C/G/T/A: 5/0/5/2. Found: wl-Pt; 5.1/0.1/5.0/1.8, w2-Pt; 5.0/0.3/4.9/1.8.
139
Pt-DNA adduct stability test. Experiments were performed to investigate whether heating
platinated oligonucleotides in thiol-containing buffer causes removal of the Pt cross-link.
Samples of w2-Pt were dissolved in either annealing buffer containing 100 mM NaCl, 70 mM
Tris/HCl pH 7.5, 10 mM MgCl 2 , 5 mM DTT, or pure water to a concentration of ~0.2 mM. Each
sample was heated to 90 'C and slowly cooled to 4 'C over 3 h, while a control sample in water
was kept at 4 'C. All samples were kept in the dark throughout the experiment. The solutions
were then dialyzed against water at 4 'C in the dark over 24 h, and Pt/DNA ratios were
determined. The control sample maintained a Pt/DNA ratio of 1.00 ± 0.01, whereas the heated
samples in water or buffer showed Pt/DNA ratios of 0.97 ± 0.01 and 0.92 ± 0.01, respectively.
Synthesis of site-specifically platinated DNA duplexes ti-Pt and t2-Pt. Ligation of the
individual oligonucleotide strands to form the 146-bp DNA duplex containing either a 1,3-cis{Pt(NH 3)2 }2+-d(GpTpG) (ti-Pt) or 1,2-cis-{Pt(NH 3)2 }2+-d(GpG) (t2-Pt) intrastrand cross-link
was performed by phosphorylation of the 5'-OH groups of the pertinent components, followed by
annealing and enzymatic ligation. These conditions, previously developed to synthesize
picomoles of material for biochemical investigations,3 4 were modified and optimized here for
preparative-scale syntheses. Oligonucleotides wl, wl-Pt, w2, w2-Pt, x, yi, and y2 were
phosphorylated with T4 polynucleotide
kinase under standard conditions using DNA
concentrations between 6-8 pM. The phosphorylated strands, containing the shorter w strand in
2-fold excess, were then combined with an equimolar quantity of v and z and annealed (~3 pM
DNA, 100 mM NaCl, 70 mM Tris/HCl pH 7.5, 10 mM MgCl 2 , 5 mM DTT) from 90 'C to 4 'C
over 3 h in a PCR thermocycler using a constant temperature gradient. Ligation was performed
in situ (50 mM NaCl, 60 mM Tris/HCl pH 7.5, 10 mM MgCl 2 , 10 mM DTT, 1.5 mM ATP, 25
140
pg/mL BSA, 10 U/ptL T4 DNA ligase) over 48 h at 16 *C. Following inactivation of the enzyme
at 65 'C for 20 min, the sample was dialyzed against water overnight at 4 'C, then lyophilized,
reconstituted in formamide, and purified by 6% urea-PAGE. DNA was extracted from the gel,
ethanol precipitated, and re-annealed in ~250 ptL buffer. Duplex DNA was purified from residual
single-stranded DNA by 6% native-PAGE; the desired product was extracted from the gel into
buffer (50 mM NaCl, 10 mM Tris/HCl, 1 mM EDTA), lyophilized to dryness, reconstituted in
water, and ethanol precipitated before quantitation by UV-Vis and AA spectroscopy. Syntheses
were performed on a 6.0 - 25.0 nmol scale, with typical overall yields of 15 - 20%.
Restriction enzyme digestion of tl-Pt/t2-Pt. Pt-DNA adducts located at a restriction site inhibit
cleavage by restriction enzymes.4 ' 43 The ti oligomer contains a single HpyCH4III recognition
sequence (5'-ACNG*T-3') at the GTG platination site, whereas the t2 oligomer contains an
EcoRI sequence (5'-G*AATTC-3') that overlaps the GG adduct target. In order to evaluate the
fraction of each DNA duplex platinated at the desired site, t1/t1-Pt and t2/t2-Pt strands were
incubated with HpyCH4III and EcoRI, respectively, and the cleavage products were analyzed by
denaturing gel electrophoresis. The DNA duplexes were 5'-labelled with
32P
on both strands,
then 10 pmol of each strand was incubated with the appropriate restriction enzyme for 1 h at 37
'C in buffer provided by the manufacturer. Samples were subsequently phenol extracted to
remove the enzyme, ethanol precipitated, dissolved in formamide, and analyzed by 6% ureaPAGE.
Purification of histone core proteins H2A, H2B. Frozen cell paste containing one of the
expressed histone proteins (from X laevis, expressed in E. coli) were thawed and suspended in
141
60 mL of wash buffer (50 mM Tris-HCl pH 7.5, 100 mm NaCl, 1 mM EDTA, 2 mM DTT) in a
stainless steel beaker. The suspension was sonicated in an ice bath for 24 min (30 sec pulses) to
lyse the cells, centrifuged at 40,000 RPM for 45 min at 4 'C, and the supernatant decanted. The
pellet was re-suspended in 60 mL Triton wash buffer (wash buffer containing 1% v/v Triton100) to solubilize the cell membrane, centrifuged, and decanted. This process was repeated
twice, followed by a final wash with buffer without surfactant. The pellet was soaked in 1 mL
DMSO for 1 h at room temperature, then suspended in 10 mL of unfolding buffer (7 M
guanidine-HCl, 20 mM Tris-HCl pH 7.5, 10 mM DDT). The mixture was incubated at 37 'C for
1 h to solubilize the protein, then centrifuged and passed through a 0.45 pm syringe filter.
The supernatant was loaded onto a pre-equilibrated Pharmacia Sephacryl S-200 26/60
high resolution gel filtration column and eluted with SAU- 1000 buffer (7 M deionized urea, 20
mM Na acetate pH 5.2, 1 M NaCl, 2 mM DTT, 1 mM EDTA) at 1 mL/min. Collected fractions
were analyzed by UV-Vis spectroscopy and SDS-PAGE. Fractions containing histone protein
were pooled and dialyzed (1000 MWCO) against three changes of 2 L of 2 mM DTT over 24 h
at 4 0 C, then lyophilized to dryness. The dried sample was dissolved in 10 mL of SAU-200
buffer (7 M deionized urea, 20 mM Na acetate pH 5.2, 0.2 M NaCl, 2 mM DTT, 1 mM EDTA)
and loaded onto a Pharmacia 5 mL Hi-Trap SP cation exchange column, pre-equilibrated with
SAU-200 buffer.
Protein was eluted at 1 mL/min, while slowly increasing the NaCl
concentration to 0.6 M over 80 min. Fractions were analyzed by UV-VIS spectroscopy and
SDS-PAGE. Those containing pure protein were combined and dialyzed (1000 MWCO) against
three changes of 2 L of 2 mM DTT over 24 h at 4 0C, then lyophilized to dryness.
142
Histone octamer refolding and purification. Approximately 3.5 mg of each of the four
lyophilized histone core proteins (H2A, H2B, H3, and H4) were dissolved in unfolding buffer (7
M guanidine/HCl, 20 mM Tris/HCl pH 7.5, 10 mM DTT) to a concentration of ~ 2 mg/mL. The
exact concentration was determined by UV-Vis spectroscopy. The proteins were allowed to
unfold for 1 h, then equimolar portions of each of the proteins were combined and the final
protein concentration adjusted to 1 mg/mL. The sample was dialyzed (6-8000 MWCO) against 3
changes of 2 L refolding buffer (2 M NaCl, 10 mM Tris/HCl pH 7.5, 1 mM EDTA, 5 mM 2mercaptoethanol) over 24 h at 4 'C. The sample was centrifuged at 3000 x g for 5 min at 4 'C to
remove precipitate, then concentrated to
-1
mL using a Centricon centrifugal concentrator. The
concentrated sample was loaded onto a pre-equilibrated Superdex Hi-Load 16/60 gel filtration
column and eluted at 0.6 mL/min with refolding buffer for 10 h. Fractions were analyzed by
UV-Vis spectroscopy and SDS-PAGE (Fig. 5.4) and those containing octamer were combined
and concentrated to 1 mL using a centrifugal filter device. The sample was diluted to 50% v/v
with glycerol, then stored at -24 'C.
H2A/H2B
H3
H4
H4
1
2
3
4
5
6
Figure 5.4. SDS-PAGE analysis of fractions for histone octamer purification. Lanes 1, 2:
histone octamer, lanes 3, 4: H2A-H2B dimer, lane 5: protein ladder, lane 6: H2A-H2B dimer.
Assembly of nucleosome core particles. Nucleosome core particles were assembled from DNA
duplexes t1-Pt or t2-Pt and histone octamers according to published procedures.4 4 Briefly,
143
histone octamer was combined with 0.8 equiv of either ti-Pt or t2-Pt in 2 M KCl, 10 mM
Tris/HCl pH 7.5, 1 mM EDTA, and 1 mM DTT in a dialysis membrane. The stoichiometry of
the assembly was determined experimentally to maximize the yield. The DNA concentration in
solution was approximately 0.7 mg/mL. The sample was placed in a dialysis vessel containing
400 mL of the same buffer at 4 'C. Over 18 h low salt buffer (0.25 M KCl, 10 mM Tris/HCl pH
7.5, 1 mM EDTA, and 1 mM DTT) was introduced into the vessel using a peristaltic pump at a
rate of 1.5 mL/min while buffer was removed at the same rate, thus keeping the total volume of
dialysis buffer constant. The sample was then dialyzed against 400 mL of low-salt buffer and
three changes of 400 mL of 20 mM Tris/HCl pH 7.5, 1 mM EDTA, and 1 mM DTT, each for 4 h
at a time. Finally, each sample was heat equilibrated at 45 'C for 2 h in order homogenize the
translational position of DNA in the nucleosomes.
The tl-Pt-NCP sample was concentrated to -100 pL and purified by preparative
electrophoresis (4.5% native PAGE, 0.3X TBE, 37.5:1 mono:bis acrylamide, 5 cm column
length, 29 mm internal diameter) using a Biorad 491 prep cell connected to an Akta FPLC pump
and fraction collector. Fractions were analyzed by 4.5% native PAGE, and samples containing
pure centrally-phased nucleosomes (lanes 5 and 6, Fig. 5.5) were combined and concentrated to
1 mg/mL with a Centricon YM-10 device. The next fraction (lane 7, Fig 5.5) was 96% pure,
calculated by quantitation of the ethidium bromide stained gel, and was isolated and concentrated
separately. Each sample was dialyzed against CCS buffer (20 mM potassium cacodylate pH 6.0,
1 mM EDTA, 1 mM DTT, three changes of 400 mL at 4 C), and further concentrated to -8
mg/mL.
144
1
2
3
4
5
6
7
8
9
10
500
350
300
250
200
Figure 5.5. Purification of tl-Pt-NCP by preparative gel electrophoresis. lane 2: unshifted
sample prior to gel purification, lane 3: shifted sample before purification, lanes 4-10: collected
fractions from prep cell. Fractions 5-6 were combined, and fraction 7 was collected separately.
Nucleosomes from t2-Pt were purified via a different protocol because the scale was
insufficient for preparative electrophoresis. After heat equilibration and concentration, the
sample was purified on 4.5% native PAGE, and the NCP bands were visualized and excised by
UV shadowing. Nucleosomes were isolated from the gel slices with a Millipore Centrilutor
system at 200 V for 2 h at 4 'C with Centricon 50K centrifugal filters, using 0.2X TBE as an
elution buffer. The buffer was exchanged with CCS buffer, and the sample concentrated to -25
pM (~ 5 mg/mL). Yields for both ti-Pt and t2-Pt nucleosome samples were determined by UVVis spectroscopy (A260) using an extinction coefficient of 10 AU = 1 mg/mL NCP. Typical
isolated yields for nucleosomes containing either ti-Pt or t2-Pt were 20-40%.
Crystallization studies. Diffraction-quality crystals of tl-Pt-NCP, containing the cisplatin 1,3{Pt(NH 3)2 }2 -d(GpTpG) intrastrand cross-link, were grown by sitting-drop vapor diffusion under
conditions described in previous nucleosome structural studies.1 5 Droplets (1 pL) containing -20
pM PtNCP, 75-80 mM MnCl 2 , 55-60 mM KCl, and 20 mM potassium cacodylate pH 6.0 were
equilibrated at 20 'C against 200 pL of precipitant solution containing 40-46 mM MnCl 2, 30-45
mM KCl, and 20 mM potassium cacodylate pH 6.0. Crystals grew in 1-2 weeks, and varied in
145
size between -10 x 40 x 100 pm and -0.2 x 0.2 x 0.5 pm. Identical conditions were explored for
the t2-Pt-NCP (containing the 1,2-{Pt(NH 3)2 }2 -d(GpG) adduct), but no crystals were obtained.
Several cryoprotecting conditions were explored in order to determine the optimal
procedure for handling tl-Pt-NCP nucleosome crystals. In all cases the final conditions were 37
mM MnCl2 , 40 mM KCl, 20 mM potassium cacodylate pH 6.0, 24% v/v 2-methyl-2,4pentanediol (MPD), and 2% w/v trehalose. Three protocols were attempted in which the
nucleosome crystals were either soaked in cryosolution overnight or soaked in cryosolution for
approximately 20 min. For the third condition, crystals were transferred to a solution of 37 mM
MnCl 2, 40 mM KCl, 20 mM potassium cacodylate pH 6.0, 4% v/v 2-methyl-2,4-pentanediol, and
2% w/v trehalose, and the MPD concentration was gradually increased in situ in 5 stepwise soaks
of -2 min each until the final MPD concentration was reached. It has been previously
documented that nucleosome crystals frozen in liquid nitrogen diffract poorly,4 5 but that handling
in liquid propane has served to alleviate this problem. There is substantial debate in the literature
over the differences in freezing rates between liquid propane and nitrogen,46 '47 as well as the
effect on diffraction data quality. Thus, crystals from each group were frozen either in liquid
nitrogen at 77 K, or in liquid propane at a temperature of 140-160 K. Liquid propane-frozen
crystals were stored in cryovials in liquid nitrogen during shipment to the synchrotron.
Data collection and processing. Diffraction data were collected at 100 K at beamline 24-ID-C
at the Advanced Photon Source (APS) at Argonne National Laboratory. Data sets were collected
at either the Se-K (0.979 A) or Pt-L3 (1.072 A) edges. Between 90-120 frames were collect for a
typical data set, with ACD = 10 per frame. An example diffraction image is shown in Fig. 5.6. A
total of 19 data sets of tl-Pt-NCP crystals were collected, which were subsequently integrated
146
and scaled with HKL2000." Crystals formed in the space group P212121. Unit cell dimensions
varied with freezing conditions, with 101 A < a < 106 A, 107 A < b : 110 A, and 172 A < c <
178 A. Crystals have a solvent content of -55%. Data collection statistics are summarized in
Table 5.1.
Table 5.1. Data collection statistics for tl-Pt-NCP crystals frozen under different
cryoconditions. Values in parentheses represent data from the highest resolution bin. TRmerge I|I-(I)|II. Cryoconditions: 1, stepwise MPD soak; 2, overnight MPD soak; 3, -20 min MPD
soak.
Data set
N 2(,y/propane()
Cyrocondition
Space group
Unit cell (A)
a
b
C
Unit cell volume (A3)
Resolution range (A)
Highest res. bin (A)
No. of reflections
Completeness (%)
Redundancy
I/a(I)
Rmee (%)t
1
propaneq)
1
P2 1212 1
2
propaneq)
2
P212121
3
propaneq)
3
P21212 1
4
5
6
N2(1>)
N2(1>
N2(1)
1
P21212 1
2
P2 12121
3
P212 121
101.8
108.0
172.5
1,896,792
102.9
109.1
173.5
1,948,079
50-5.1
102.1
108.6
173.0
1,917,647
50-4.4
101.5
108.8
172.3
1,902,713
50-5.6
106.1
109.4
177,0
2,054,284
50-3.9
5.39 - 5.20
5.28
5.10
4.56 - 4.40
5.80 - 5.60
4.04
7622
98.1 (99.9)
3.7
13.5 (2.3)
10.5 (69.9)
7750
92.7 (95.8)
4.5
14.0 (2.0)
10.5 (74.3)
12629
98.3 (99.9)
3.4
21.8 (2.3)
10.7 (69.0)
4588
73.2 (76.7)
2.8
11.8 (2.3)
9.9 (65.2)
18892
98.0 (99.6)
3.7
18.8 (2.5)
7.9 (66.7)
103.8
108.8
174.1
1,964,518
50-4.0
4.14-4.00
16866
97.8 (98.9)
3.6
8.8 (1.5)
11.7 (57.2)
50
-
5.2
-
-
3.90
The data were initially processed to 3.9 A resolution. However, inspection of the image
in Fig. 5.6 reveals that diffraction is severely anisotropic, with limits to 3.2 A in the b* direction
but only 4.1 A in the c* direction. Thus the data output from HKL2000 was further truncated
using a centered ellipsoid with vertices of 1/3.6 A, 1/3.2 A, and 1/4.1 A along a*, b* , and c*,
respectively, along with anisotropic scale factor correction as described previously. 49 The data
set is 98.6% complete at 3.9 A resolution, but only 67.8% complete at 3.2 A. However,
ellipsoidal truncation allows over 3000 addition well-measured reflections to be included in the
data set, an ~15% increase, and it is critical, particularly at low resolution, to maximize the data-
147
to-parameter ratio in X-ray structure determination. A comparison of the two processed data sets
is shown in Table 5.2.
direclior
3.0
c dreclion
F/sigra
30.
10 1 7 2 58 5 1 4 5 4 1 38 3,6 3.4 3,2
resoluiomn
Figure 5.6. (left) Diffraction image of tl-Pt-NCP crystal. (right) Signal-to-noise ratios (F/s) of
diffraction data in the a*, b*, and c* directions.
Table 5.2. Comparison of diffraction data sets truncated spherically at 3.9 A, or ellipsoidally
between 3.2 and 4.1 A.
Data truncation
Space group
Unit cell (A)
a
b
c
ellipsoid
P212 121
sphere
P21212 1
106.1
109.4
177.0
106.1
109.4
177.0
Resolution range (A)
50 - 3.22
50
Highest res. bin (A)
No. of reflections
Completeness (%)
3.30-3.22
22367
67.8
4.04-3.90
18892
98.6
-
3.9
Model refinement. The 2.8 A resolution structure of the nucleosome (1AOI)
5
was used as a
search model for phasing the Pt-nucleosome data by molecular replacement using the program
Phaser. 0 Subsequent rounds of refinement and manual model building without the platinum
adduct were performed using CNS 5 1 and Coot, 52 respectively. A simulated-annealing composite
148
omit map was calculated with CNS to account for model bias. After fitting the DNA and histone
core structure to the composite omit map density, the platinum adduct was inserted into the
appropriate location (vide infra). Final refinement of the Pt-NCP model was performed in
Refmac5, with manual model adjustments carried out in Coot. Temperature factor refinement
was attempted using grouped B-factors as described, individual isotropic B-factors, and a pureTLS model5 3,54 using the (H3/H4) 2 tetramer, two H2A/H2B dimers, and each DNA strand as
separate TLS groups. TLS refinement provided the best agreement between the model and the
data, as reflected by the Rfree value. Final refinement statistics are given in Table 5.3.
Table 5.3. Model refinement statistics.
Resolution range (A)
50 - 3.90
24.9
Rwork(%)'
Rfree(%)'
30.6
B-factors (A2)
Protein
DNA
RMSD bond lengths
RMSD bond angles
Protein atoms
DNA atoms
tRmerge
=
101.5
219.9
0.013 A
1.600
6051
5980
ElfIN/ EI.
= I|F0\ - |Fe|||/JFo|.
R value obtained for a test set of reflections (5% of diffraction data).
R
Results
Synthesis of site-specifically platinated mononucleosomes. Synthetic 146 bp DNA containing
a centrally engineered 1,3-cis-{Pt(NH 3)2 }2+-d(GpTpG) or 1,2-cis-{Pt(NH 3)2}2+-d(GpG) crosslink was prepared and purified on a milligram scale by enzymatic ligation of five component
oligonucleotides. Atomic absorption spectroscopy, mass spectrometry, and restriction enzyme
digestion confirmed the presence of one platinum atom per DNA duplex. Pt/DNA ratios for the
wl-Pt or w2-Pt starting material ranged from 1.03 to 1.15, and MALDI mass spectra (data not
149
shown) gave no evidence for the presence of multiply platinated species. The slightly higher
Pt/DNA ratios are most likely due to inaccurate extinction coefficients for UV absorbance at 260
nm, which were calculated based on base composition and do include a contribution from the
platinum adduct. Pt/DNA ratios for ti-Pt or t2-Pt ranged from 0.85 to 0.92. Digestion of the ti,
ti-Pt, t2, or t2-Pt strands with either HpyCH4III (for GTG-containing strands) or EcoRI (for
GG-containing strands) provided evidence that the final construct retains the Pt lesion (see Fig.
5.7). One sample was contaminated with a significant fraction of unplatinated DNA and was
discarded; all other samples were shown to contain >90% pure platinated DNA, as determined
by quantitation of the autoradiographed gel. It appears, based on Pt/DNA ratios calculated before
and after ligation, that a small fraction of platinum is removed from DNA during synthesis.
Polynucleotide kinase and T4 DNA ligase buffers contain 5 mM and 10 mM concentrations of
DTT, respectively, and it is possible that excessive heating or prolonged time in solution may
allow the sulfur-containing species to coordinate to platinum and remove the adduct from DNA.
To test this hypothesis, Pt/DNA ratios on w2-Pt were calculated before and after subjecting the
sample to annealing conditions at 90'C in PNK buffer. The Pt/DNA ratio decreased from 1.00
0.01 to 0.92 + 0.01 after heating. These results may be identical within experimental error, but
they possibly indicate that a small fraction of the cisplatin cross-link is removed during DNA
ligation.
Nucleosomes were prepared in high purity from platinated DNA and recombinant histone
octamers by dialysis. Several hundred micrograms of tl-Pt-NCP were obtained after preparative
electrophoresis, which yields a product of higher purity compared to the electroelution method,
provided that the synthesis is performed on a scale greater than -200 pg. Although several
advancements in both DNA ligation efficiency and nucleosome reconstitution and purification
150
improved the overall yield of platinated nucleosomes, synthesizing sufficient quantities of PtDNA was a significant challenge for this project. A total of ~0.5 mg of tl-Pt-NCP and 32 pg of
t2-Pt-NCP were isolated for crystallization studies.
146-GG
146-GTG
Pt
Digest
-
+
-
+
-
+
+
+
+
-
+
+
-
+
+
+
-
-
-
+
+
+
250
4
200
150
0
100
75
50
Figure 5.7. Restriction enzyme digestion of t1, ti-Pt, t2, or t2-Pt. Platinated DNA blocks
digestion by either HpyCH4III or EcoRI. Bands at ~75 nt in the Pt-containing samples indicate
unplatinated impurities.
Crystallization and data collection. Crystals of tl-Pt-NCP, containing the 1,3-d(GpTpG)
cisplatin adduct, grew readily under the conditions described in other nucleosome structure
papers.55 '56 The size of these crystals varied significantly, but generally they were smaller than
reported sizes in other nucleosome crystallization experiments. Macroseeding attempts did not
yield larger crystals, but these crystals were still sufficiently large to be analyzed using the MD-2
microdiffractometer beam at APS beamline 24-ID-C, operated by the Northeast Collaborative
Access Team (NE-CAT), which is capable of focusing to a 10 jim radius. Unfortunately crystals
of t2-Pt-NCP were never obtained. The reasons for this failure are unclear, but insufficient
material was obtained to perform a full crystallization investigation, and the primary researcher
on this project (Paresh Agarwal) left the laboratory before the work could be completed.
151
A thorough study of the effects of cryofreezing temperature on diffraction quality
demonstrated that NCP crystals should be frozen in liquid propane at a temperature of-153 K to
maximize data quality and resolution.4 5 Direct freezing in liquid nitrogen led to contraction of
the unit cell along the c axis from 181 A to 175 A and significant loss of resolution. Several
different cryosoaking procedures have been reported in the literature that include (i) a multistep
soak in increasing concentrations of MPD, (ii) a short, direct soak, or (iii) overnight incubation
of the crystal in cryosolution. For diffraction analysis of tl-Pt-NCP crystals, samples were
frozen both in liquid nitrogen of liquid propane, and all three cryoprotection techniques were
used. However, in this study no advantage was gained by cryoprotecting crystals at higher
temperature. The highest diffracting propane-frozen crystals showed reflections to 4.4 - 5.2 A,
whereas nitrogen-frozen crystals diffracted to 3.9 - 5.6 A (see Table 5.1).
Structure of the platinated nucleosome.
The nucleosome accommodates
a 1,3-cis-
{Pt(NH 3)2 }2 +-d(GpTpG) cross-link without any significant affect on the overall DNA or protein
structure (see Fig. 5.8). N-terminal tails of each histone are disordered and not visible in the
structure, as is the case in almost all other nucleosome models, and the octamer core structure
does not deviate from other X-ray structures of nucleosomes incorporating X laevis histones.
DNA adopts the same conformation around the protein core as undamaged nucleosomes, with
hydrogen-bonding interactions between primarily arginine and lysine residues and the DNA
phosphodiester backbone stabilizing the complex. These contacts occur at fourteen locations
along the duplex, each time the minor groove reaches the protein core. In addition the DNA in
this model and the original crystal structure determination5 5 maintain the same rotational phasing
(shown in Fig. 5.9) because the platinum cross-link was specifically incorporated at a location on
152
the DNA strand where the nucleosome rotational position would match that of the native
nucleosome core particle. In this way the chances of crystallizing the complex under similar
conditions would be maximized.
(A)
a
(D)
Figure 5.8. The structure of the platinum-damaged nucleosome core particle. (A) Overall NCP
structure, which closely matches that of native nucleosomes. (H3/H4) 2 tetramer is shown in
green, and H2A/H2B dimers in blue. (B) Top view of the 1,3-cis-{Pt(NH 3)2}2 -d(GTG) crosslink (in purple/yellow). The dyad axis is marked by (D.(C) and (D) 2F-Fe electron density maps
surrounding the platinated DNA segment and an H3 a-helix, respectively.
The histone proteins are more ordered in the structure than the DNA superhelix (compare
electron density maps in Fig. 5.8c and d)., as evidenced by the average temperature factors. Bfactors for all protein and DNA atoms averaged 101.5 A2 and 219.9 A2 , respectively. The most
ordered DNA bases are those that contact the histone core, and the least ordered are those facing
the solvent, leading to a periodic distribution of temperature factors that is exactly out of phase
153
for the two DNA strands (see Fig. 5.10). Watson-Crick base-pairing was restrained during
refinement, which aided the modeling of DNA regions facing the solvent having unclear electron
density. Although the DNA backbone, particularly the phosphate groups, and overall double
helical structure are discernible (see Fig. 5.8c), individual base pairs are not resolved in the
electron density, so geometric parameters could not be accurately determined.
1
73 %L74
146
Pt(GTG)
Figure 5.9. Overlay of platinated nucleosomal DNA (in blue) with DNA from the original
nucleosome structure (red). The platinated sequence is shown in cyan. All crystallographic
images were created with Pymol. 57
400
350
300
250
200
150
100
50
20
40
60
80
100
120
140
Base pair
Figure 5.10. B-factor distribution of phosphorus atoms of the DNA backbone. Blue line
represents the platinated strand (5' -+3'), and red line the unplatinated strand (3' -+5').
154
Because of the pseudo symmetry of nucleosome core particles about the dyad rotational
axis, 55 NCPs can pack into the crystal lattice in either of two possible orientations. Many X-ray
crystal structures incorporate a palindromic DNA sequence in order to circumvent this potential
problem. In the present structure both the DNA sequence and position of the 1,3-cis{Pt(NH 3)2 }2+-d(GpTpG) adduct are asymmetric with respect to the 2-fold symmetry axis, so two
possible orientations could be present. An asymmetric nucleosome core particle has previously
been studied by X-ray crystallography, but the resolution (3.2
A) is notably lower than almost all
other structures determined by the research group. The resolution of the present structure made
location of the platinum adduct in initial refinement stages ambiguous, and the DNA bases were
not individually resolved. However, since the 1,3-cis-{Pt(NH 3)2 }2+-d(GpTpG) cross-link was
site-specifically engineered in the DNA duplex, placement of the adduct was limited to two
possible positions.
In nucleosome core particles containing 146 bp DNA, 1 base pair falls directly on the
dyad axis, splitting the DNA into "long" and "short" halves containing 73 and 72 bp segments,
respectively.55 Three NCP models were therefore prepared during refinement, in which the
cisplatin 1,3-intrastrand cross-link was placed on either the short or long DNA segment, each of
its two possible locations on either side of the pseudo-symmetry dyad axis, or in which no
platinum moiety was included. Each model was refined through identical procedures using CNS
with grouped B-factor minimization (2 groups per residue); later refinements were subsequently
performed in Refmac5. The Rfree values for models without platinum, or with platinum on either
the short or long DNA segment, were 30.9%, 30.6%, and 32.3%, respectively, indicating
superiority for the orientation in which the 1,3-cis-{Pt(NH 3)2}2+-d(GpTpG) adduct is located on
the 72 bp DNA half. This model and the one without platinum contained the same DNA
155
sequence, the difference being only the presence or absence of a {Pt(NH 3 )2}2 + moiety. Because
the entire DNA duplex in the two platinum-containing models was oriented differently, the gap
in Rfree values between the resulting structures was more dramatic. An F-Fe electron density
difference map calculated from the model lacking any cisplatin adduct contained a positive peak
at the appropriate position on the short DNA segment, but not at the other possible location,
providing additional evidence that the proper orientation was chosen (Fig. 5.11). Furthermore, a
difference map calculated from the model containing the cisplatin adduct in the incorrect
orientation on the long DNA half gave rise to a large negative peak surrounding the platinum
atom. Together these data confirm that the cisplatin intrastrand cross-link location is properly
determined in the reported nucleosome structure. Location of the platinum atom was attempted
by calculating anomalous difference Fourier maps using data collected at 1.072 A, but the single
platinum atom provided insufficient anomalous signal for heavy atom location.
1
2
2
Figure 5.11. Possible 1,3-cis-{Pt(NH3)2 }2+-d(GpTpG) cross-link locations on the nucleosome
core particle. Position 1 is located on the 72 bp DNA half of the dyad axis (marked by <D), and
position 2 is located on the long half (73 bp). (1) F-Fe electron density map (green) calculated
from a structure with the platinum atom omitted, contoured at 5y, indicates the correct platinum
position. (2) F-Fe electron density map (in red) contoured at -50 reveals incorrect placement of
the Pt cross-link.
156
The 1,3-cis-{Pt(NH 3)2 }2+-d(GpTpG) cross-link faces inward toward the octamer and
away from the solvent exposed surface. This DNA rotational setting agrees with that predicted
by chemical footprinting experiments (see Fig.5.12),. No clear interactions between the cisplatin
lesion and octamer core are observed in the structure. One of the platinum ammine ligands sits
-5.0 A from a lysine side-chain amino group, and these moieties may interact by water-mediated
hydrogen-bonding contacts, which is commonly observed at the DNA-histone surface. 58 Details
of the 1,3-cis-{Pt(NH 3)2}2 +-d(GpTpG) adduct geometry also could not be discerned from the
electron density, so Pt-N bond distances and angles were restrained during model refinement to
values typical for platinum(II) square planar coordination compounds. In particular, the extruded
thymine base is disordered, as it is in NMR solution structures of the platinated DNA. 6
3' dG
3 dG
S'dG
'_
6'dG
O CTAME R
"
OCTAM ER
Figure 5.12. Stereo view of the 1,3-cis-{Pt(NH 3)2}2 +-d(GpTpG) cross-link, looking down the
DNA double helix. The location of the histone octamer is marked, showing how the cisplatin
intrastrand adduct faces inwards towards the protein core.
Discussion
The X-ray crystal structure of a nucleosome core particle modified with a specifically
engineered 1,3-cis-{Pt(NH3)2} 2+-d(GpTpG)cross-link reveals interesting details about the effects
of cisplatin-DNA damage on nucleosome structure. The Pt intrastrand cross-link is positioned
near the dyad axis and faces the histone octamer core, in agreement with previous solution
157
studies.3"'
This orientation has implications for DNA repair, because the adduct may be
"hidden" from nucleotide excision repair machinery and inhibit removal. Repair of Pt-DNA
adducts is less efficient on nucleosomal compared to free DNA.
Interestingly, DNA near the platinum cross-link adopts a conformation similar to that of
free DNA having a centrally located 1,3-Pt(GpTpG) adduct, as determined in solution by NMR
spectroscopy. 6 An 11-bp DNA segment encompassing the Pt-DNA adduct (4 bp on each side of
the cross-link) was compared to the free DNA NMR solution structure containing the 1,3-cis{Pt(NH 3)2 }2 -d(GpTpG) cross-link (see Fig. 5.13) The helical bend angles for the nucleosomal
and free DNA segments are 39.10 and 45.4', respectively, indicating that the local nucleic acid
structure around the Pt adduct mimics its solution-state form. Values were calculated using the
program Curves+, with the NMR value deviating from that reported in the original publication.
This discrepancy arises from known and controversial differences in calculating global DNA
structure parameters.59
These results suggest a mechanism by which cisplatin intrastrand cross-links might direct
nucleosomal DNA to a specific rotational position that accommodates the structural deviations
caused by the bifunctional adduct. Superhelical DNA in the nucleosome is highly distorted, and
the data presented here indicate that Pt-DNA adducts alter the DNA position in the nucleosome
such that the bend induced by a platinum 1,3-intrastrand cross-link is congruent with the bend
caused by wrapping of DNA around the histone core. Further evidence to support this conclusion
is provided by the observation that nucleosome core particles treated with cisplatin or oxaliplatin
form DNA adducts preferentially at locations where the purine bases already experience a large
roll angle due to the superhelical structure. 3 6 Inspection of base pair parameters in the highresolution nucleosome X-ray structure (Fig. 5.14) reveals that the roll angle, which varies
158
periodically along the superhelix, is maximized at locations on the DNA equivalent to the
relative position of the cisplatin adduct in the present structure. Lack of discernible electron
density on individual base pairs prevents such an analysis here.
X-ray (NCP)
Bend angle = 46.40
NMR (DNA)
Bend angle = 39.10
Figure 5.13. Global helical bend angle of the NMR solution structure of duplex DNA containing
the 1,3-cis-{Pt(NH 3)2}2 -d(GTG) cross-link (left), 6 and a DNA segment containing the adduct
taken from the nucleosome structure (right). Gray lines represent the helical axis, calculated with
Curves+.
Further analysis of the cisplatin-modified nucleosome core particle is limited by a low
resolution limit and insufficient electron density in sections of the DNA and platinum intrastrand
cross-link. Several factors probably contribute to the relative DNA disorder. First, the exterior
folds of macromolecular structures are typically less ordered than the interior sections. Second,
all nucleosome structures exhibit higher average DNA temperature factors for DNA atoms than
for protein atoms, suggesting that the nucleic acid strands generally experience higher thermal
motion. The refined temperature factors from a pure-TLS model in this structure are very high,
but comparable to those obtained for other nucleosome core particles at similar resolution.36 6 0
Finally, because the DNA sequence is non-palindromic and the Pt-DNA adduct is asymmetric
with respect to the nucleosome dyad, orientation of NCPs within the crystal lattice is a potential
159
problem. That the platinum B-factor is larger than that of the local surrounding phosphate
backbone is further evidence that orientational ambiguity contributes to the DNA disorder, and
possibly the overall resolution limit. An asymmetric nucleosome structure has been solved
previously,6 0 but notably, the resolution of this structure was limited to 3.2 A when the majority
of NCP structures from this group are at 2.0 - 2.5 A resolution. These deficiencies prevent a
more detailed analysis of the DNA structure. However the presented model in which the cisplatin
cross-link is placed in the current position is clearly superior to the alternate conformation, as
indicated by the Rfee values of each model and Fo-Fe electron density maps.
GAATC,A'iGAACAG
20
u
WGAGTTCAAATACACTTTGGTAGTATCTGCAGGTGGATATT
1TTGAT
-r~-
EIJ-
ago nna
s
- Hri
oA
a-
o
n
MA &to
a
n
10
-10
a -20
Figure 5.14. Analysis of roll angle of the DNA superhelix in the high-resolution X-ray crystal
structure of the nucleosome core particle (PDB accession code: 1KX5, shown in red). Locations
along the DNA where roll angle is maximized (marked by yellow arrow and yellow DNA) aligns
with placement of the platinum cross-link in this structure (blue, with platinum cross-link
location shown in yellow).
Conclusions
Nucleosome core particles containing either a 1,2-cis-{Pt(NH 3)2}2+-d(GpG) or 1,3-cis{Pt(NH 3)2 }2+-d(GpTpG) intrastrand cross-link located near the dyad axis were synthesized, and
160
the X-ray crystal structure of latter construct was solved at 3.2 A resolution. The overall structure
is conserved between Pt-damaged and unmodified nucleosomes, and the platinum lesion, thought
to be the major adduct formed by carboplatin-DNA binding, adopts a conformation pointing
inward toward the histone core which is consistent with previous biochemical experiments. The
DNA structure in the vicinity of the platinum intrastrand cross-link is similar to that in solution
as determined by NMR spectroscopy. These results indicate that Pt cross-links direct rotational
phasing of nucleosomal DNA to a position where the structural changes caused by platinum
binding are readily accommodated by the nucleosomal superhelix. The current model accurately
mimics the environment of cisplatin-DNA damage in cancer cells by incorporating the
nucleosome structure found in all eukaryotic cells.
161
References
(1)
Jamieson, E. R.; Lippard, S. J. Chem. Rev. 1999, 99, 2467-2498.
(2)
Wang, D.; Lippard, S. J. Nat. Rev. Drug Discov. 2005, 4, 307-320.
(3)
Todd, R. C.; Lippard, S. J. Metallomics 2009, 1, 280-29 1.
(4)
Takahara, P. M.; Frederick, C. A.; Lippard, S. J. J. Am. Chem. Soc. 1996, 118, 1230912321.
(5)
Gelasco, A.; Lippard, S. J. Biochemistry 1998, 37, 9230-9239.
(6)
Teuben, J.-M.; Bauer, C.; Wang, A. H.-J.; Reedijk, J. Biochemistry 1999, 38, 1230512312.
(7)
Wu, Y.; Bhattacharyya, D.; King, C. L.; Baskerville-Abraham, I.; Huh, S.-H.; Boysen,
G.; Swenberg, J. A.; Temple, B.; Campbell, S. L.; Chaney, S. G. Biochemistry 2007, 46,
6477-6487.
(8)
Coste, F.; Malinge, J.-M.; Serre, L.; Shepard, W.; Roth, M.; Leng, M.; Zelwer, C. Nucleic
Acids Res. 1999, 27, 1837-1846.
(9)
Lovejoy, K. S.; Todd, R. C.; Zhang, S.; McCormick, M. S.; D'Aquino, J. A.; Reardon, J.
T.; Sancar, A.; Giacomini, K. M.; Lippard, S. J. Proc. Natl. Acad. Sci. U.S.A. 2008, 105,
8902-8907.
(10)
Komberg, R. D.; Lorch, Y. Cell 1999, 98, 285-294.
(11)
Workman, J. L. Genes Dev. 2006, 20, 2009-2017.
(12)
Segal, E.; Fondufe-Mittendorf, Y.; Chen, L.; Thistr6m, A.; Field, Y.; Moore, I. K.;
Wang, J.-P. Z.; Widom, J. Nature 2006, 442, 772-778.
(13)
Muthurajan, U. M.; Park, Y.-J.; Edayathumangalam, R. S.; Suto, R. K.; Chakravarthy, S.;
Dyer, P. M.; Luger, K. Biopolymers 2003, 68, 547-556.
(14)
Richmond, T. J.; Davey, C. A. Nature 2003, 423, 145-150.
(15)
Luger, K.; Mader, A. W.; Richmond, R. K.; Sargent, D. F.; Richmond, T. J. Nature 1997,
389, 251-260.
(16)
White, C. L.; Suto, R. K.; Luger, K. EMBO J2001, 20, 5207-5218.
(17)
Clapier, C. R.; Chakravarthy, S.; Petosa, C.; Fem-ndez-Tomero, C.; Luger, K.; M,1ler, C.
W. Proteins:Structure, Function, andBioinformatics 2008, 71, 1-7.
(18)
Tsunaka, Y.; Kajimura, N.; Tate, S.-i.; Morikawa, K. Nucl. Acids Res. 2005, 33, 34243434.
162
(19)
Luger, K. Chromosome Res. 2006, 14, 5-16.
(20)
Pennings, S.; Meersseman, G.; Bradbury, E. M. J. Mol. Biol. 1991, 220, 101-110.
(21)
Flaus, A.; Owen-Hughes, T. Curr. Opin. Genet. Dev. 2001, 11, 148-154.
(22)
Park, Y.-J.; Chodaparambil, J. V.; Bao, Y.; McBryant, S. J.; Luger, K. J. Biol. Chem.
2005, 280, 1817-1825.
(23)
Saha, A.; Wittmeyer, J.; Cairns, B. R. Genes Dev. 2002, 16, 2120-2124.
(24)
Reinberg, D.; Sims III, R. J. J. Biol. Chem. 2006, 281, 23297-23301.
(25)
Lippard, S. J.; Hoeschele, J. D. Proc.Natl. Acad Sci. U.S.A. 1979, 76, 6091-6095.
(26)
Simpkins, H.; Pearlman, L. F. FEBS Lett 1984, 169, 30-34.
(27)
Hayes, J. J.; Scovell, W. M. Biochem. Biophys. Acta 1990, 1088, 413-418.
(28)
Galea, A. M.; Murray, V. Biochem. Biophys. Acta 2002, 1579, 142-152.
(29)
Marzilli, L. G.; Reily, M. D.; Heyl, B. L.; McMurray, C. T.; Wilson, W. D. FEBS Lett
1984, 176, 389-392.
(30)
Koyama, Y.; Kikuchi, S.; Nakagawa, S.; Kobayashi, S. Chem. Pharm. Bull. 2007, 55,
520-525.
(31)
Danford, A. J.; Wang, D.; Wang,
U.S.A. 2005, 102, 12311-12316.
(32)
Ober, M.; Lippard, S. J. J. Am. Chem. Soc. 2007, 129, 6278-6286.
(33)
Ober, M.; Lippard, S. J. J Am. Chem. Soc. 2008, 130, 2851-2861.
(34)
Wang, D.; Hara, R.; Singh, G.; Sancar, A.; Lippard, S. J. Biochemistry 2003, 42, 67476753.
(35)
Wu, B.; Davey, C. A. Chem. Biol. 2008, 15, 1023-1028.
(36)
Wu, B.; Droge, P.; Davey, C. A. Nat. Chem. Biol. 2008, 4, 110-112.
(37)
Davies, N. P.; Hardman, L. C.; Murray, V. Nucleic Acids Res. 2000, 28, 2954-295 8.
(38)
Blommaert, F. A.; van Dijk-Knijnenburg, H. C. M.; Dijt, F. J.; den Engelse, L.; Baan, R.
A.; Berends, F.; Fichtinger-Schepman, A. M. J. Biochemistry 1995, 34, 8474-8480.
(39)
Dhara, S. C. Indian J. Chem. 1970, 8, 193-194.
(40)
Kibbe, W. A. Nucl. Acids Res. 2007, 35, W43-W46.
Q.; Tullius,
163
T. D.; Lippard, S. J. Proc.Natl. Acad. Sci.
(41)
Todd, R. C.; Lovejoy, K. S.; Lippard, S. J. J. Am. Chem. Soc. 2007, 129, 6370-6371.
(42)
Cohen, G. L.; Ledner, J. A.; Bauer, W. R.; Ushay, H. M.; Caravana, C.; Lippard, S. J. J.
Am. Chem. Soc. 1980, 102, 2487-2488.
(43)
Ang, W. H.; Myint, M. N. Z.; Lippard, S. J. J. Am. Chem. Soc. 2010, submitted.
(44)
Dyer, P. N.; Edayathumangalam, R. S.; White, C. L.; Bao, Y.; Chakravarthy, S.;
Muthurajan, U. M.; Luger, K. Methods Enzymol. 2004, 375, 23-44.
(45)
Edayathumangalam, R. S.; Luger, K. Acta Crystallogr.2005, D61, 891-898.
(46)
Teng, T.-Y.; Moffat, K. JournalofApplied Crystallography1998, 31, 252-257.
(47)
Walker, L. J.; Moreno, P. 0.; Hope, H. J. Appl. Cryst. 1998, 31, 954-956.
(48)
Otwinowski, Z.; Minor, W. Methods Enzymol. 1997, 276, 307-326.
(49)
Strong, M.; Sawaya, M. R.; Wang, S.; Phillips, M.; Cascio, D.; Eisenberg, D. Proc. Natl.
Acad. Sci. US.A. 2006, 103, 8060-8065.
(50)
McCoy, A. J.; Grosse-Kunstleve, R. W.; Adams, P. D.; Winn, M. D.; Storoni, L. C.;
Read, R. J. J Appl. Cryst. 2007, 40, 658-674.
(51)
BrUnger, A. T.; Adams, P. D.; Clore, G. M.; DeLano, W. L.; Gros, P.; Grosse-Kunstleve,
R. W.; Jiang, J.-S.; Kuszewski, J.; Nilges, M.; Pannu, N. S.; Read, R. J.; Rice, L. M.;
Simonson, T.; Warren, G. L. A cta Crystallogr.1998, D54, 905-92 1.
(52)
Emsley, P.; Cowtan, K. Acta Crystallogr.2004, D60, 2126-2132.
(53)
Winn, M. D.; Isupov, M. N.; Murshudov, G. N. Acta Crystallogr.2001, D57, 122-133.
(54)
Winn, M. D.; Murshudov, G. N.; Papiz, M. Z. Methods Enzymol. 2003, 374, 300-32 1.
(55)
Luger, K.; Mader, A. W.; Richmond, R. K.; Sargent, D. F.; Richmond, T. J. Nature 1997,
389, 251-260.
(56)
Rhodes, D.; Brown, R.; Klug, A. Methods Enzymol. 1989, 170.
(57)
DeLano, W. L. DeLano Scientific, PaloAlto, CA, USA 2002.
(58)
Davey, C. A.; Sargent, D. F.; Luger, K.; Maeder, A. W.; Richmond, T. J. J. Mol. Biol.
2002, 319, 1097-1113.
(59)
Lavery, R.; Moakher, M.; Maddocks, J. H.; Petkeviciute, D.; Zakrzewska, K. Nucleic
Acids Res. 2009, 37, 5917-5929.
(60)
Bao, Y.; White, C. L.; Luger, K. J. Mol. Biol. 2006, 361, 617-624.
164
Chapter 6. Exploring Transcription by T7 RNA Polymerase from Free and Nucleosomal
DNA Modified with Site-Specific Platinum Intrastrand Cross-links
165
Introduction
Studies involving both bacteriophage T7 RNA polymerase (T7 RNAP) 1 5 and eukaryotic
RNA polymerase III (RNAP 111)6 have demonstrated that nucleosomal DNA can be transcribed
without removal of the histone octamer; however, the rate of RNA synthesis is significantly
lower than on free DNA. Bacteriophage polymerases do not encounter nucleosomes in vivo, but
they can be used as a model to study transcription by RNAP III, because both enzymes operate
by similar mechanisms, depicted in Fig. 6.1.6,7 The transcription elongation complex initially
disrupts DNA-histone contacts -20 base pairs ahead of the polymerase. As the complex reaches
the nucleosome dyad, the octamer is displaced to a DNA region behind the RNA polymerase.
During this process an intermediate loop forms, which is transcribed slowly and is perceived to
be the rate-limiting step. After transfer of the histone core behind the elongation complex is
completed, transcription resumes at a high rate. Advancement of the polymerase along
nucleosomal DNA induces rotational strain in the double helix, which is released through a twist
diffusion mechanism. This mechanism is thought to be critical to the fidelity of nucleosome
transcription. 8, A series of DNA-binding pyrrole-imidazole ligands inhibit nucleic acid twist
propagation, and also block transcription by T7 RNAP from nucleosomal DNA, but not from
free DNA. These results indicate a direct correlation between functional nucleosome mobility
and successful transcription.
In contrast to T7 RNAP or RNAP III, RNA polymerase 1I (RNAP 1I) is strongly inhibited
by nucleosome barriers at physiological salt concentrations. At higher ionic strength in vitro, this
barrier is relieved and transcription resumes concomitant with release of an H2A/H2B dimer
from the histone core. 10-12 Transcription by RNAP II in vivo occurs by a different mechanism,
facilitated by ATP-dependent chromatin remodeling complexes
166
3
that dissociate nucleosomes
downstream of the elongation complex to remove the barrier (see Fig. 6.1).14,1 The mechanism
of this process, however, is not completely understood.
a)
b)
H2A-H28
po ode9
re-
odeng
L
§
2A/H28
Figure 6.1. Mechanisms of nucleosomal transcription. (a) T7 RNA polymerase or RNA
polymerase III can transcribe through nucleosomes by histone translocation. (b) Eukaryotic RNA
polymerase II requires chromatin remodeling complexes to remove the histone barrier to
transcription.
Inhibition of transcription by platinum-DNA damage has been investigated thoroughly,
as reviewed elsewhere;1 6 however, many of these studies were performed in vitro and utilized
linear DNA that do not account for nucleosome structure. Biochemical experiments17-19 and Xray crystal structure evidence described in Chapter 5 demonstrate that cisplatin-DNA crosslinks direct nucleosomal DNA toward a specific rotational phasing that overrides the native
influence of even strong nucleosome positioning sequences. Other data indicate that globally
platinated DNA inhibits histone translocation and propagation of twist diffusion in a manner
similar to that for the aforementioned pyrrole-imidazole complexes. 20 These data suggest a
167
mechanism whereby cisplatin-DNA adducts may inhibit transcription by denying RNA
polymerase elongation complexes access to nucleosomal DNA. However, the ratio of
administered platinum per DNA base pair used in these experiments was too high to be
physiologically relevant, and global platination of nucleosomes does not permit the contribution
of a specific platinum intrastrand cross-link to be evaluated, which is important in mechanistic
investigations. Therefore more research is necessary to test the validity of this hypothesis.
This chapter describes experiments performed to explore (i) whether a single 1,2-cis{Pt(NH 3)2 }2 +-d(GpG) or 1,3-cis-{Pt(NH 3)2 }2+-d(GpTpG) intrastrand cross-link located near the
nucleosome dyad will inhibit DNA twist propagation, and (ii) how elongation complexes of T7
RNAP navigate nucleosomes modified with site-specific cisplatin intrastrand adducts on either
the template or coding strands. The former was measured by examining ATP-independent
nucleosome mobility, which requires propagation of rotational strain along the nucleosomal
DNA.2
Heating of nucleosome samples causes off-centered, kinetically formed NCPs to
equilibrate to thermodynamically favored centered positions through histone translocation. The
ratio of each complex can be determined by native gel electrophoresis, which can resolve these
species. To investigate the latter objective, DNA or nucleosome templates were synthesized and
subjected to single-round transcription assays by T7 RNAP. Nucleosomes containing centrallylocated cisplatin 1,2-d(GpG) or 1,3-d(GpTpG) intrastrand adducts on either the template or
coding strands were assembled from 145 base pair DNA with a 9 nucleotide overhang and
recombinant histones. These constructs were ligated to a biotinylated 50 bp DNA fragment
containing a T7 RNAP promoter and complementary overhang, and immobilized on
streptavidin-coated magnetic beads. Assembling the nucleosomes initially on shorter DNA
ensures a uniform translational position of the histone octamer on the transcription template.
168
Nucleosome positions were assessed by restriction enzyme mapping of the DNA strand. The
template DNA sequence lacks adenine between the promoter and ligation sites, so that
elongation complexes can be directed past the ligation site by incubation with nucleotide
triphosphates lacking UTP. This step ensures that only transcription of ligated templates will be
recorded. Transcription stalls at the first adenine base, and resumes upon addition of the full set
of NTPs, including a
32 P-labelled
nucleotide. Resulting RNA transcripts were resolved by gel
electrophoresis.
Experimental
Materials. Phosphoramidites, columns, and other reagents for solid-phase oligonucleotide
synthesis were purchased from Glen Research. Potassium tetrachloroplatinate, which was used to
synthesize cisplatin,2 2 was a gift from Engelhard Corporation (now BASF). Enzymes were
32
purchased from New England Biolabs. y- P-ATP and a-3 2P-GTP (6000 Ci/mmol) were
purchased from Perkin Elmer. Dynabead M-280 streptavidin-coated magnetic beads were
purchased from Invitrogen. All other reagents were obtained from commercial suppliers and
used without further purification. UV-Vis spectra were recorded with a Hewlett-Packard 8453
instrument. Liquid chromatography was performed with an Agilent 1200 series HPLC equipped
with a temperature-controlled autosampler and automated fraction collector. All dialyses
employed Spectra/Por dialysis membranes of an appropriate molecular weight cut-off, and were
pre-treated with hot 50 mM EDTA, followed by several washes with water and dialysis buffer,
prior to use. Atomic absorption spectroscopy, used to quantitate platinum concentrations, was
performed with a Perkin Elmer AAnalyst 300 system. Radioactive gels were visualized using a
Storm 840 phosphorimager, and radioactivity in samples was determined with a Beckman LS
169
6500 scintillation counter. All nucleosome gels were run on 4.5% native PAGE (0.3X TBE,
37.5:1 mono:bis acrylamide) in a cold room at 4C.
Preparation of s1/s2/s1-Pt/s2-Pt duplexes. DNA duplexes containing either a cisplatin 1,3d(GpTpG) (si-Pt) or 1,2-d(GpG) (s2-Pt) adduct on the template DNA strand were synthesized
as previously described.' 8 Oligonucleotides depicted in Fig. 6.2 were either prepared in-house
using an Applied Biosystems Model 392 DNA/RNA synthesizer or purchased from Integrated
DNA Technologies and purified by denaturing gel electrophoresis prior to use. Platinated strands
dl-Pt and d2-Pt were prepared by reaction of cisplatin with dl or d2, respectively, in 10 mM
HEPES pH 6.8 buffer as described previously,2 3 and purified by ion-exchange HPLC using
method RCT.006S (Appendix C). HPLC chromatograms of the purification of each strand are
shown in Figs. 6.3 and 6.4. Pt/DNA ratios for dl-Pt and d2-Pt, calculated by atomic absorption
and UV-Vis spectroscopy, respectively, were 1.05 ± 0.13 and 1.17 ± 0.02, respectively.
Individual strands were ligated to form the 154/145 bp DNA duplex as depicted in Fig.
6.2b. Oligonucleotides a, bi, b2, c, dl, dl-Pt, d2, d2-Pt, and g were phosphorylated with T4
polynucleotide kinase under standard conditions at DNA concentrations of 6 ptM. Annealing of
duplexes si, s2, si-Pt, and s2-Pt (0.8 pM DNA, 100 mM NaCl, 70 mM Tris/HCl pH 7.5, 10 mM
MgCl 2 , 5 mM DTT) and ligation (50 mM NaCl, 60 mM Tris/HCl pH 7.5, 10 mM MgCl 2 , 10 mM
DTT, 1.5 mM ATP, 25 ptg/mL BSA, 10 U/pL T4 DNA ligase) were performed as described in
Chapter 5. The 145mer and 154mer single strands were resolved and isolated separately by 5%
denaturing PAGE. Equimolar portions of each strand were re-annealed for each duplex, and
dsDNA was purified by 6% native-PAGE. The desired product was extracted from the gel into
buffer (50 mM NaCl, 10 mM Tris/HCl, 1 mM EDTA), ethanol precipitated, reconstituted in
170
water, and quantitated by UV-Vis and AA spectroscopy. Syntheses were performed on a 1 nmol
scale; yields for the isolated single strand DNA ranged from 38 - 65%. Pt/DNA ratios for si-Pt
and s2-Pt were 0.93 ± 0.03 and 0.95 ± 0.02, respectively.
(A)
a (7Omer):
5'TAAATTAATAGTTGAAGTTGTAGTAAATGTTAATGTAGATCTGTTGTTCCGATATTACCAAAA
CCTTCAC
hl (75mer):
5'CTAATAGCGCTAGTACACAGGAGAAGGACATGAACATGAACCTAATGAACACAACAAATAATG
TAAGTGCCCATG
b2 (75mer):
5'CTAATAGCGCTAGTAACCAGGAGAAGGACATGAACATGAACCTAATGAACACAACAAATAATG
TAAGTGCCCATG
c (91mer):
5'TAGCGCTATTAGGTGAAGGTTTTGGTAATATCGGAACAACAGATCTACATTAACATTTACTAC
AACTTCAACTATTAATTTACCCAGTGCC
dl (14mer):
5'CTTCTCCTGTGTAC
dl-Pt (Pt-14mer):
5'CTTCTCCTGTGTAC
d2 (14mer):
5'CTTCTCCTGGTTAC
d2-Pt (Pt-14mer):
5'CTTCTCCTGGTTAC
e (49mer):
5'CATGGGCACTTACATTATTTGTTGTGTTCATTAGGTTCATGTTCATGTC
f (59mer):
5'TAATACGACTCACTATAGGGAGCCGACAACACCCGGAGCCGACAACACCCGGCACTGGG
g (50mer):
5'GGGTGTTGTCGGCTCCGGGTGTTGTCGGCTCCCTATAGTGAGTCGTATTA
bio-g (3'-biotion-50mer):
5'GGGTGTTGTCGGCTCCGGGTGTTGTCGGCTCCCTATAGTGAGTCGTATTA-biotin
(B)
70-a
91-c
1 ligation
7514-Pt
145
49-.
154-Pt
2 NCP assemby
594
3.fgsbon__
171
zz
Figure 6.2. (on previous page) (a) Oligonucleotides synthesized for preparation of nucleosomal
DNA containing single cisplatin cross-links on the template strand. Bases shown in red depict
platination sites. (b) Reaction scheme showing ligation of oligonucleotides a-e to form a 145 bp
duplex with 9 nt overhang, assembly of nucleosome core particles, and ligation of a biotinylated
(purple sphere) 50 bp fragment containing a T7 RNAP promoter site to form the full 204 bp
template.
300
250
200
150
100
50
0
5
10
20
15
25
30
35
40
45
50
Time (min)
Figure 6.3. HPLC purification of dl-Pt (peak 1) from unplatinated dl (peak 2) and DNA strands
containing multiple platinum atoms bound (marked by
60
50
40
30
20
10
0
0
5
10
15
25
20
30
35
40
Time (min)
Figure 6.4. HPLC purification of d2-Pt (peak 1) from unplatinated d2 (peak 2) and DNA strands
containing multiple platinum atoms bound (marked by *).
Preparation of c1/c2/c1-Pt/c2-Pt duplexes. DNA duplexes with cisplatin cross-link sites (cl-Pt
=
1,3-cis-{Pt(NH 3)2}2+-d(GpTpG), c2-Pt
=
1,2-cis-{Pt(NH 3)2}2+-d(GpG)) on the 145 nt, non-
172
template strand were synthesized in the same manner, as described above. The oligonucleotides
in Fig. 6.5 were utilized for these syntheses to form the double-stranded constructs. The 14mer
strands for cisplatin binding were of the same sequence as those used in s duplex synthesis, in
order to conserve material. The only other change in DNA sequence from that in the s strands
was the abolition of a native T7 RNAP termination sequence, ATCTGTT, which caused
premature release of the polymerase elongation complex (vide infra). The four duplexes c1, c2,
cl-Pt, and c2-Pt were prepared on a 1.2 nmol scale in yields ranging from 48 to 65%.
(A)
a (82mer):
TAAATTAATAGTTGAAGTTGTAGTAAATGTTAATGTAGATGTGATGTTCCGATATTACCAAAACC
TTCACCTAATAGCGCTA
dl (14mer):
5'CTTCTCCTGTGTAC
dl-Pt (Pt-14mer):
5'CTTCTCCTGTGTAC
d2 (14mer):
5'CTTCTCCTGGTTAC
d2-Pt (Pt-14mer):
5'CTTCTCCTGGTTAC
c (49mer):
GACATGAACATGAACCTAATGAACACAACAAATAATGTAAGTGCCCATG
bl (75mer):
CATGGGCACTTACATTATTTGTTGTGTTCATTAGGTTCATGTTCATGTCGTACACAGGAGAAGTA
GCGCTATTAG
b2 (75mer):
CATGGGCACTTACATTATTTGTTGTGTTCATTAGGTTCATGTTCATGTCGTAACCAGGAGAAGTA
GCGCTATTAG
e (79mer):
GTGAAGGTTTTGGTAATATCGGAACATCACATCTACATTAACATTTACTACAACTTCAACTATTA
ATTTACCCAGTGCC
(B)
82
79
14(Pt)
49
145(P)
75
154
173
Figure 6.5. (on previous page) (a) Oligonucleotides synthesized for preparation of nucleosomal
DNA containing single cisplatin cross-links on the coding strand (c duplexes). Bases shown in
red depict platination sites. (b) ligation scheme to form c1/c2 duplexes.
Assembly and purification of nucleosome core particles. Nucleosome core particles were
assembled from eight DNA duplexes: si, si-Pt, s2, s2-Pt, c1, cl-Pt, c2, and c2-Pt. A 40 pmol
portion of DNA and 50 pmol of the recombinant histone octamer were combined in 70 piL of
reconstitution buffer according to published procedures.
Purification of histone proteins and
folding of the octamer core were described in Chapter 5. After dialysis, each sample was heat
equilibrated at 45 'C for 2 h. A 2 pmol aliquot of each sample was analyzed by 4.5% native
PAGE and stained with ethidium bromide (see Fig. 6.6). Initially, nucleosome samples were
purified by native PAGE; NCP bands were visualized by UV shadowing and excised from the
gel. Nucleosome core particles were isolated from the gel by electroelution with a Millipore
Centrilutor system into Centricon YM-10 centrifugal concentrators (0.3X TBE, 200 V, 2 h at
4C). Each sample was concentrated to 100 pL, then the buffer was exchanged with 20 mM
Tris/HCl pH 7.5, 1 mM EDTA, 1 mM DTT by centrifugation. The concentration of nucleosomes
was determined by UV-Vis spectroscopy using an extinction coefficient of 10 AU = 1 mg/mL
NCP. Yields for s1-NCP, s2-NCP, si-Pt-NCP, and s2-Pt-NCP ranged from 60-70 pmol (43 50%). However, subsequent electrophoretic analysis of purified samples revealed that a minor
(15-20%) amount of DNA contamination persisted, presumably due to the known effect that
dilute nucleosome solutions cause dissociation of DNA from the histone octamer." Subsequent
sample sets were prepared and used without further purification. The amount of free DNA in
each sample was quantitated from the ethidium bromide-stained gel.
174
Seq:
Pt:
s1
S1
s2
s2
C1
Cl
c2
c2
-
+
-
+
-
+
-
+
-NCP
-
DNA
Figure 6.6. Assembly of nucleosome core particles of 145 bp DNA duplexes, as shown by native
PAGE. Gel was visualized by ethidium bromide staining.
Nucleosome mobility investigation. Nucleosomes, assembled from either ti, t1-Pt, or t2-Pt
DNA containing no platinum, a cisplatin 1,3-d(GpTpG) intrastrand cross-link, or a 1,2-d(GpG)
adduct, respectively, (sequence and synthesis described in Chapter 5) were investigated to
determine the effect of the Pt-DNA adduct on ATP-independent nucleosomal mobility.
32p_
Labelled tI, ti-Pt, or t2-Pt strands (40 pmol) were combined with an equimolar amount of
histone cores in 20 ptL of buffer containing 2 M KCl, and nucleosomes were prepared by dialysis
as described.2 4 All samples were prepared in duplicate. Following dialysis, the volume of each
sample was brought to 50 pL, and a 20 pL portion of each was incubated at 37 *C or 50 'C. The
remaining 10 pL was kept at 4 'C. Aliquots of each sample were taken at 30, 60, 120, and 180
min, and the radioactivity was quantified by scintillation counting. Samples were analyzed by
4.5% native PAGE.
Preparation of 204 bp immobilized transcription templates
A biotinylated and radiolabelled promoter fragment containing the T7 RNA polymerase
promoter site was prepared by combining 100 pmol each of 5'-32 P-labelled f and 5'phosphorylated bio-g (sequences in Fig. 6.2), annealing from 80 'C to 4 'C over 2.5 h using a
175
constant temperature gradient, and purifying by 5% native PAGE. For transcription experiments,
unlabelled DNA was utilized. The promoter fragment (2 pmol) was ligated with an equimolar
portion of either DNA or NCP with sequence si, si-Pt, s2-Pt, ci, cl-Pt, or c2-Pt in 100 gL
solution (50 mM Tris/HCl pH 7.5, 10 mM MgCl 2 , 1 mM ATP, 10 mM DTT, 0.5 mg/mL BSA,
1%PEG-8000, and 0.4 U/ptL T4 DNA ligase) at 16 'C for 2 h. Samples were heated to 50 'C for
10 min to deactivate the enzyme. Separately, 1.5 mg of Dynabead M-280 streptavidin-coated
magnetic beads were washed two times each with 200 piL of 0.1 M NaOH, 50 mM NaCl in
diethylpyrocarbonate (DEPC)-treated water, then 100 mM NaCl in DEPC-treated water to make
suitable for RNA applications. Beads were then washed two times each with bind/wash buffer (2
M NaCl, 10 mM Tris/HCl pH 7.5, 1 mM EDTA) and TE600 buffer (600 mM NaCl, 10 mM
Tris/HCl pH 7.5, 1 mM EDTA), then resuspended in 750 p.L of TE600. Washing was performed
by suspending the sample by pipette mixing, then collecting the beads with a magnet and
removing the supernatant. After deactivation of the T4 DNA ligase, a 100 pL aliquot of
streptavidin beads was added to each sample, mixed by pipette, and incubated at room
temperature for 30 min with gentle rocking to allow binding of the biotinylated DNA constructs.
Unbound material was removed by washing the beads three times with TE300, and three times
with transcription buffer (40 mM Tris-HCl pH 7.9, 6 mM MgCl 2 , 2 mM spermidine, 10 mM
NaCl, 10 mM DTT, in nuclease-free water). For transcription experiments, the beads were
resuspended in 200 ptL of transcription buffer for future use. For analysis of the ligation
products, samples were incubated in 20 pL of 95% formamide, 25 mM EDTA at 90 'C for 5 min
to remove biotinylated DNA from the streptavidin constructs, and loaded directly onto a 6%
urea-polyacrylamide gel for analysis (Fig. 6. 7).
176
NCP
DNA
1
204 nt
product
59 nt
2
3
1
2
3
-*
1*W
*oO
promoter
Figure 6.7. Gel analysis of ligation to form 204-bp transcription templates with free or
nucleosomal DNA. Lanes 1 = s1, 2 = si-Pt, 3 = s2-Pt.
Restriction enzyme mapping of nucleosome position
Restriction enzyme digestions of ligated templates were performed in order to assess the
position of nucleosome core particles on the 204 bp construct after ligation. Streptavidin-bound
transcription templates were prepared with s2-Pt radiolabelled with
32
P at the 5' end of the
template strand with and without the nucleosome according to the procedure described above,
washed, and resuspended in TE buffer. Aliquots of each of the ligated products were then
subjected to digestion with SfcI, BsrI, BglII, HaeII, or Bsp12861, each of which have a single
cutting site along the DNA strand, as shown in Fig. 6.8. In the general reaction, 1.0 pmol of 204
bp DNA or NCP was incubated with 1 pL of restriction enzyme in 100 pL of commercially
supplied buffer for 1 h at 37 'C. As control reactions, unligated s2-Pt DNA or NCP was digested
with BsrI, BglII, HaeII, or Bsp12861 in a similar manner. Enzymes and histones were removed
177
from solution by phenol extraction, which also releases biotinylated DNA from the magnetic
beads, and product in the supernatant was ethanol precipitated twice, dissolved in formamide,
and analyzed by 5% urea-PAGE.
Ski
(13117)
owtBgll
(5&157)
(968100)
Heell
(139/135)
Bep12861
(200/196)
Figure 6.8. Restriction enzyme sites along the 204-bp DNA. The blue oval represents DNA
covered by the nucleosome. (A/B) refer to the 5' and 3' terminal base, respectively. The
radiolabel was located on the 5' end of the bottom DNA strand.
Single-round in vitro transcription assays with T7 RNA polymerase
Single-round promoter-dependent transcription by T7 RNA polymerase was performed
based on two previously reported procedures, as shown in Fig. 6.9.26,27 Immobilized DNA or
NCP templates were pre-equilibrated with transcription buffer after ligation, as described in the
previous section. Initial transcription walking past the promoter ligation site was carried out in
20 ptL of transcription buffer containing 25 nM DNA or NCP, 10 U T7 RNA polymerase, 1.0
U/pL RNasin (RNase inhibitor), 25 pM ATP, 25 pM GTP, and 25 ptM CTP at room temperature
for 5 min. Transcription stalls after synthesis of a 37 nt RNA transcript due to the lack of UTP in
solution. The supernatant was removed, and the immobilized transcription elongation complexes
were washed five times each with 100 pL reaction buffer. Radiolabeling of the RNA transcript
was achieved by incubating the washed elongation complexes in 20 pL transcription buffer
containing 1.0 U/ptL RNasin, 1.0 pM UTP, and 0.6 pM a-32P-GTP at room temperature for 5
min. This step allows the polymerase to incorporate the next five nucleotides, including three
178
32 P-labelled
GMP units, and stalls after synthesis of a 42 nt transcript. Finally, transcription is
completed by addition of 0.5 mM NTPs to the solution and incubation for 15 min at room
temperature, allowing the polymerase to transcribe off the template or release at any point along
the DNA. Unlabelled GTP is present in 1000-fold excess in the final reaction solution, so
subsequent rounds of transcription incorporate non-radiolabelled nucleotides and only the first
round of RNA synthesis is visualized on a gel. The reaction was quenched by addition of an
equal volume of 20 mM EDTA, and the supernatant was ethanol precipitated, dissolved in
formamide, and analyzed by 6% urea-PAGE. For kinetic analysis, transcription was performed at
0 'C, and aliquots of the final transcription solution were taken at 0, 15, 45, and 90 s for DNA
templates, and at 0, 15, 30, 60, and 90 s for nucleosome templates.
inte
l~tanrprnsrpiteate
C-;!tedsideprouct
1) + ATP, GTP, CTP
"Transcription walking" beyond ligation site
2) + UTP, 1-P-GTP
Radiolabelling of RNA transcript
3) + all 4 NTPs
complete transcription
..
.-
.
.
179
Figure 6.9. (on previous page) Experimental system using immobilized free or nucleosomal
templates to study transcription by T7 RNA polymerase. Elongation complexes are formed on a
mixture of fully ligated templates and unligated promoter strands. The polymerase is directed
past the ligation site by adding ATP, GTP, and CTP. After washing away the NTPs, RNA
transcripts are radiolabelled by adding a- 32 P-GTP, then transcription is completed by addition of
all four nucleotide triphosphates. RNA transcripts (dashed line) are analyzed by denaturing
PAGE.
Results and Discussion
Preparation of immobilized, site-specifically
platinated free and nucleosomal DNA
transcription templates. Synthesis of 145 bp DNA engineered with either a 1,3-cis{Pt(NH 3)2 }2+-d(GpTpG) or 1,2-cis-{Pt(NH 3)2}2+-d(GpG) cisplatin intrastrand cross-link and a 9
nt overhang was achieved, and nucleosome core particles were assembled from these constructs
and recombinant histone proteins from X laevis in high yield. Ligation yields for the s1, s2, c1,
or c2 duplexes were significantly higher (-45% overall) than for t1 or t2 duplexes (-15%)
described in Chapter 5, despite both samples having been prepared under similar reaction
conditions. One possible reason for this discrepancy is that t1/t2 duplexes were synthesized on a
much larger scale, 10-20 nmol compared to -1 nmol syntheses of transcription DNAs, and
isolation protocols were not fully optimized for larger scale reactions. In particular, extraction of
oligonucleotides from gels after purification becomes problematic when the quantity of
polyacrylamide is increased. Typical isolation techniques, such as the use of centrifugal filtration
or syringe filters, lead to clogging and result in lost material. Another possibility is that because
t1/t2 DNA is divided in two -70 nt halves with very similar nucleotide sequences, there can be
annealing issues. Ligation of the full duplex requires that the five starting oligonucleotides
combine to form a specific double helix; any undesired duplex combination would lower the
overall ligation yield. Sequences of the sl/s2 DNA components are less self-complementary
compared to those of the former, so enzymatic ligation yields increase as a consequence.
180
Nucleosomes were assembled from all eight 145 bp DNA duplexes by gradient dialysis
and isolated by native gel electrophoresis and electroelution in good yield. However, at low
concentration in solution, DNA will dissociate from the histone octamer and introduce a free
DNA contaminant in the sample.2 s Thus nucleosome preparations were used for transcription
without further purification, and remaining histone octamer in the sample was washed from the
beads after ligation of the full template. Final transcription constructs were synthesized by
ligating 145 bp DNA/NCP and the biotinylated 50 bp T7 RNAP promoter-containing fragment
and immobilizing the biotinylated product on streptavidin-coated magnetic beads. The 204-bp
strand is the only ligation product. Ligation of free DNA proceeds more efficiently than ligation
of nucleosomal DNA, as demonstrated by the relative amounts of product and starting material
(Fig. 6.7). This effect is probably due to physical interactions between the histone core and T4
DNA ligase.
Restriction enzyme mapping (shown in Fig. 6.10) was utilized to assess the nucleosome
position after formation and immobilization of the full template. The cutting efficiency of each
enzyme is nearly 100% on free DNA, as shown by digestion of either s2-Pt or the full DNA
transcription template (lanes 2-6 and 11-16, respectively, in Fig. 6.10). BsrI does not cleave s2Pt DNA because the restriction site is located on the 9-nt overhang (lane 3), but it cleaves the
204 bp construct after ligation of the overhang (lane 13).
The s2-Pt nucleosome sample contains 86% nucleosomal DNA and 14% residual free
DNA, based on quantitation of analytical PAGE (data not shown). The amount of undigested
DNA for BglII, HaeII, and Bsp 12861 (lanes 8-10), the restriction sites of which are covered by
the histone octamer, vary between 50-70%, indicating that these enzymes are partially able to
recognize the restriction site and cleave nucleosomal DNA. After ligation the BsrI site is cut
181
almost entirely (lane 18), indicating that the restriction site is exposed and nucleosome position
does not change after synthesis of the longer DNA construct. However, a higher level of
digestion by BglII, HaeII, and Bsp12861 is observed after ligation and immobilization on
streptavidin-coated beads (lanes 19-21), indicating that some dissociation of histones from the
DNA occurs. The amounts of undigested DNA in the s2-Pt nucleosome or full nucleosome
template in each sample are compared in Table 6.1, to approximate the amount and location of
nucleosomes after ligation of the promoter site, binding to the solid support, and washing the full
template. Based on differences in restriction enzyme digestion, it is estimated that this procedure
results in 15-20% loss of octamer from the DNA. Although histone dissociation is not optimal, it
is more important to have a uniformly positioned nucleosome core particle on the template in
order to examine the mechanism of transcription along NCPs, because the amount of free DNA
in each sample can be measured and corrected for in subsequent experiments. As a consequence
of residual free DNA left over from nucleosome assembly and dissociation of the histone
octamer during promoter ligation and immobilization steps, -40% of DNA templates are nonnucleosomal.
Table 6.1. Restriction enzyme mapping of nucleosome core particle position. Values represent
fraction of undigested DNA compared to total radioactivity in sample. Ligated nucleosomes are
more sensitive to digestion by BglII, HaeII, and Bsp12861 compared to unligated nucleosomes,
indicating some dissociation of histone octamer from DNA during ligation.
SfcI
BsrI
BglII
HaeII
Bsp12861
s2-Pt NCP
n/a
1.0
0.57
0.70
0.50
182
204 bp NCP
0.31
0.05
0.47
0.50
0.22
McP:
--
Ligate:
Digest:
-
--
--
+-
-
1
2
3
4
1
2
3
Lan.: 1 2
3
4
5
6
7
8
9 10 11 12 13 14 15 16 17 18 19 20 21 22
--
-
*+ .4
4
-
1
+
2
+
3
4
+
5
-
1 2
3
4
5
250
204 (full transcription template)
200 4W
187 (SfcI)
150
100 4
___
.PIP
0
-
154 (unligated s2-Pt)
147 (BsrI)
104 (BglII)
75 0
SM
*
69
(HaeII)
8S
(Bsp12861)
25S
Figure 6.10. Gel analysis of restriction enzyme mapping of nucleosome core particle position.
Nucleotide length and the corresponding restriction enzyme are shown on the right. Lanes 2-6:
s2-Pt DNA, lanes 7-10: s2-Pt NCP, lanes 11-16: 204-bp DNA, lanes 17-22: 204 bp NCP.
Digestion enzymes are 1: BsrI, 2: BglII, 3: HaeII, 4: Bsp12861, 5:SfcI.
Nucleosome mobility investigation. The ability of a single cisplatin intrastrand cross-link to
inhibit ATP-independent, heat-induced nucleosome mobility was explored. NCPs were prepared
from t1, ti-Pt, or t2-Pt DNA containing no platinum, a 1,3-cis-{Pt(NH 3)2}2+-d(GpTpG) crosslink, or a 1,2-cis-{Pt(NH 3)2}2+-d(GpG) intrastrand adduct, respectively, and subjected to heat
equilibration at either 37 'C or 50 'C for a determined time (see Fig. 6.11). At 37 'C, both
cisplatin intrastrand cross-links inhibited DNA translocation to some degree (see graph in Fig.
6.12). Unplatinated nucleosomes were completely shifted within 30 min at 37 'C, whereas
nucleosomes containing the l,3-cis-{Pt(NH 3)2}2+-d(GpTpG) cross-link required 120 min to
183
equilibrate fully at the same temperature. NCPs modified with the 1,2-Pt(GpG) intrastrand
adduct still contained -10% of nucleosomes on the off-centered translational position after 3
hours of heat equilibration, demonstrating that 1,2-Pt(GpG) cross-links have a greater inhibitory
effect than the 1,3-Pt(GpTpG) counterpart. At 50 'C all nucleosome samples were equilibrated
completely within 30 min, indicating that the mechanism of inhibition of Pt-DNA cross-links
does not involve covalent interactions between the platinum lesion and protein core, or any other
irreversible phenomenon. These results are consistent with previous work demonstrating that
nucleosome core particles globally treated with either cisplatin or oxaliplatin exhibit decreased
nucleosome mobility.2 8
off-centered
NCP
Heat tine (min) 0
1,2-d(GG)
30 60 120 18
0
centered
4
_NCP
1,3-d(GTG)
30 60 120 180 0
no Pt
30 60 120 180
Heat
time
(mn) 0
Wet-
YMP
HMW -
HMW-
off-centered
NCP- &MAoff-centered
centered
NCP - 40--
DNA-
'.0
-
-
*A
1,3-d(GTG)
30 0 120
180 0
no Pt
30 80 120 180
NCP
centered
NCP
-
-
1.2-d(GG)
30
0 120 180 0
DNA-
00
-
Figure 6.11. Native PAGE analysis of nucleosome mobility investigation of platinated
nucleosome core particles at 37 'C (left) or 50 'C (right). The off-centered NCP, a kinetic
product, converts to the more thermodynamically stable centered nucleosome conformation.
These results further demonstrate that, like nucleosomal DNA-binding polyamide
ligands, intrastrand cross-links from cisplatin inhibit DNA translocation along the histone
octamer. 8 Polyamides block transcription of nucleosome DNA but not of linear DNA, leading to
the hypothesis that these compounds inhibit transcription by locking the nucleosome in place and
184
preventing DNA sliding. This process is proposed to occur through inhibition of DNA twist
diffusion; rotation of the DNA by one-half turn would change the location of a polyamide
binding site relative to the histone octamer, and a bound ligand could prevent this translocation.
Similarly, cisplatin-DNA cross-links have both a highly preferred relative location on
nucleosomal DNA, where the adduct faces inwards toward the core, and a propensity to inhibit
thermal translocation. The preference for cisplatin cross-links to face inward toward the histone
core is discussed in Chapter 5; Pt-DNA adducts may direct DNA rotational phasing such that
DNA bend aligns with the bend of superhelical DNA in the nucleosome. Translocation of the
histone core along platinum-modified DNA would force bent Pt-DNA region out of phase with
the superhelical bend, which is disfavored. The cisplatin 1,2-d(GpG) cross-links cause a more
dramatic bend angle than 1,3-d(GpTpG) cross-links, 29 ,30 and are a stronger inhibitor of DNA
sliding. However, the magnitude of this effect from a single cisplatin adduct is smaller than that
from both the pyrrole-imidazole ligands8 and multiple Pt lesions.2 8 Therefore it cannot be
concluded from these data that a single Pt cross-link will inhibit transcription via this mechanism
analogously to polyamide ligands.
35%
50%
30%
40%
-+-1,2-GG
-9- 1,3-GTG
-r-un-Pt
* 30%
20%
15%
10%
1,2-GG
20%5%
--
1,3-GTG
10%
-*-
un-Pt
5%
0%
0%
0
50
100
150
200
Time (min)
0
50
100
150
200
Time (min)
Figure 6.12. Quantitation of nucleosome mobility of platinated samples at either 37 'C (left) or
50 'C (right). Plots show conversion of kinetic NCPs to the thermodynamically-preferred
centered nucleosomes. Error bars represent the range of values observed.
185
Single-round in vitro transcription assays. Similarities between the abilities of cisplatin
intrastrand cross-links and minor groove-binding polyamide ligands to inhibit nucleosome
sliding fuel the hypothesis that, like pyrrole-imidazole complexes, Pt-DNA adducts may block
RNA synthesis by denying polymerase access to nucleosomal DNA and stalling the elongation
complex at the histone octamer barrier. This hypothesis was tested by single-round transcription
assays of immobilized templates containing a defined nucleosome core particle and site-specific
cisplatin damage site. Transcription proceeded in three steps: (i) walking the RNA polymerase
along an A-less DNA track past the promoter ligation site using a subset of NTPs lacking UTP,
(ii) incorporation of
32
P-GMP into RNA transcripts, and (iii) completion of single-round
transcription with a full set of NTPs so that the RNA polymerase either runs off the linear
template or stalls at a defined location along the DNA.
Transcription results for constructs containing platinum on the template and coding
strands are given in Fig. 6.13. Kinetic analysis of transcription from s1 or s2 duplexes is shown
in Fig. 6.14. Three primary products are observed after transcription of DNA containing either a
1,3-cis-{Pt(NH 3)2 }2+-d(GpTpG) cross-link, or a 1,2-cis-{Pt(NH 3)2 }2+-d(GpG) intrastrand adduct,
or no platinum damage on the template strand: the 186 nt run-off transcript, a 124 nt product
resulting from polymerase stalling at the site of the platinum adduct, and a third truncated
product that appears at -90 nt in all samples (see Fig. 6.13). Transcription efficiency was
measured by comparing the relative amounts of 124 nt terminated transcript and 186 nt run-off
transcript, as shown in Fig. 6.15. Both the 1,3-d(GpTpG) and 1,2-d(GpG) cisplatin intrastrand
cross-links strongly inhibit the T7 RNAP elongation complex at the site of the cross-link. The
1,3-cross-link is measurably more effective at blocking the enzyme than the 1,2-adduct. This
trend has been observed on free DNA previously with T7 RNA polymerase in similar systems. 26
186
The native termination sequence was discovered to be the result of a T7 RNAP termination
sequence, 5'-ATCTGTT-3' on the non-template strand, a known inhibitor of T7 RNA
polymerase.
M
Coding
DNA
NCP
1 2 3 1 2 3
4-
204 nt
186 nt run-off --
\
transcript
Figure 6.13. Transcription by T7 RNA polymerase of 204-bp templates containing free or
nucleosomal DNA containing no platinum adduct (1), a 1,3-cis-Pt(GpTpG) cross-link (2), or a
1,2-cis-Pt(GpG) cross-link (3) on either the template (left) or coding strand (right). The oval
represents area of the DNA covered by the nucleosome. Pt represents the location of the crosslink, and * the location of a native termination sequence.
187
no Pt
free DNA
Pt(GTG)
Pt(GG)
no Pt
nucleosomal DNA
Pt(GTG)
Pt(GG)
Time
186 nt
run-off
42 nt
stalled
Figure 6.14. Kinetics of transcription by T7 RNA polymerase of 204-bp templates containing
free or nucleosomal DNA containing no platinum adduct, a 1,3-cis-Pt(GpTpG) cross-link, or a
1,2-cis-Pt(GpG) cross-link on the template strand. Samples were taken at 0, 15, 45, and 90 s for
DNA templates, and at 0, 15, 30, 60, and 90 s for nucleosome templates. The oval represents
area of the DNA covered by the nucleosome. Pt represents the location of the cross-link, and *
the location of a native termination sequence.
1.20c 1.00-
DNA
nucleosome
0.80-
? 0.600 S0.40-
U-
0.200.004no Pt
1,3-Pt(GTG)
1,2-Pt(GG)
Figure 6.15. Quantitation of transcription inhibition by T7 RNA polymerase from site-specific
1,3-cis-{Pt(NH3)2 }2+-d(GpTpG) or 1,2-cis-{Pt(NH 3)2 }2+-d(GpG) cross-links. Blue bars represent
transcription of DNA templates, red bars represent transcription from nucleosomal templates.
188
The same pattern of transcription inhibition was observed in free and nucleosomal DNA
samples (compare blue and red bars, Fig. 6.15). Both intrastrand cross-links nearly completely
inhibit transcription by T7 RNA polymerase when located on the DNA template strand, and the
1,3-cis-{Pt(NH 3)2}2+-d(GpTpG) cross-link
provides a stronger block than its d(GpG)
counterpart. No shorter termination sequences arising from failure of the polymerase to navigate
the nucleosome template were observed (see Fig. 6.14). This result suggests that, although a
single cisplatin intrastrand cross-link can inhibit DNA translocation along the histone octamer
required for transcription of nucleosomal DNA by T7 RNAP, the elongation complex is able to
overcome this barrier. The physical adduct, however, is a much harder obstacle to overcome, and
the polymerase is unable to transcribe through a platinum lesion placed on the template strand.
Kinetic analyses of transcription (Fig. 6.14) of both free and nucleosomal DNA demonstrate
that, under the conditions of this assay, no difference in rate of initial transcription is observed
between nucleosome templates with or without a cisplatin damage site. The bacterial RNA
polymerase can effectively transcribe through nucleosomal DNA even when the energy barrier to
nucleosome translocation is increased.
Transcription of immobilized constructs with platinum cross-links on the non-template
DNA strand yielded two major products, run-off transcript and a longer template-sized product
of 204 nt. Small amounts of RNA product were observed around 130 nucleotides in length; these
appear to arise from native termination sites because they are present in both sample sets, but the
amount varied between experiments. No transcripts corresponding to inhibition at the site of the
platinum adduct was observed because the cross-link was located on the non-template strand.
The native termination site was also abolished, confirming that the 5'-ATCTGTT-3' site was
responsible for release of the polymerase.31 Finally, no transcripts arising from stalling of the
189
elongation complex at the nucleosome barrier were observed. If platinum adducts blocked
transcription through nucleosomes by prohibiting DNA twist diffusion and translocation around
the histone core, then cross-links on both the template and coding strands would be equally
effective at restricting access to nucleosomal DNA by the RNA polymerase. These data provide
additional evidence that the enzyme can overcome both the nucleosome barrier and the
additional translocation barrier caused by platinum intrastrand cross-links. An interesting sideproduct in the transcription reactions containing platinated cross-links on the DNA coding strand
of either free or nucleosomal templates is a transcript longer than the run-off product, -204
nucleotides in length. This length corresponds to the full length of the DNA template, including
the promoter site, and is not observed in either unplatinated samples or sample containing Pt
adducts on the template strand. T7 RNAP transcripts longer than the run-off length have been
observed previously. Origins for these products include untemplated RNA synthesis,3 2
polymerase slippage along the template,3 3 3 4 RNA-templated RNA synthesis, 3 5 ,3 6 or the presence
of a DNA 3' overhang that allows the polymerase to continue transcribing the non-template
strand.3 7 3 8 Untemplated synthesis typically only adds an additional one or two nucleotides to the
RNA strand, and polymerase slippage produces a smeared band on the gel arising from a range
of transcript sizes, so these mechanisms are unlikely to be in play. RNA replication is also not
likely to occur because the transcript sequences are identical with the exception of the 3-base
sequence at the platination site, so longer transcripts would be found in unplatinated samples as
well. Template-sized RNA side products have been observed previously from DNA constructs
bearing 3' overhangs, 38 but all templates in these experiments have blunt ends. At this moment it
is unclear why a longer RNA product appears in transcription reactions utilizing DNA templates
with platinum cross-links on the coding strand.
190
The first chapter of the thesis introduces three current hypotheses describing how
platinum intrastrand cross-links may inhibit transcription in cancer cells. They may sequester
transcription factors and prevent transcription initiation, form a physical impediment around
which the elongation complex cannot navigate, or disrupt chromatin organization such that
access to nucleosomal DNA by the RNA polymerase is blocked. The results presented here
support the mechanism whereby Pt-DNA adducts located on the template strand prohibit passage
of the transcription elongation complex. Although a single {Pt(NH 3)2}2+ intrastrand cross-link
reduces the rate of nucleosome mobility to some degree, this effect is insufficient to prevent
histone translocation that occurs during transcription of nucleosomes by T7 RNA polymerase or,
by analogy, the eukaryotic RNA polymerase III. The consequences of decreased NCP mobility
by Pt-DNA damage on transcription by RNA polymerase II has not yet been investigated;
however, it seems unlikely that platinum adducts on DNA would inhibit chromatin remodeling
enzymes, which utilize ATP, that take part in the transcription process with pol II. These results
argue against disruption of nucleosome dynamics being a potential transcription inhibition
mechanism for cisplatin and other platinum antitumor drugs, and provide further evidence to
support the hypothesis that DNA adducts of these compounds physically prevent translocation of
the RNA polymerase elongation complex, even in a eukaryotic nucleosome environment.
Conclusions
The effect of a single engineered platinum intrastrand cross-link on ATP-independent
nucleosome mobility was investigated in vitro. Both 1,2-d(GpG) and 1,3-d(GpTpG) adducts of
cisplatin inhibit translocation of DNA along the histone octamer, with the former Pt lesion
providing a larger barrier. In vitro transcription assays with T7 RNA polymerase were conducted
191
to determine whether cisplatin-DNA cross-links inhibit RNA synthesis from nucleosomes
through blockage of DNA twist diffusion. Synthesis of 204 bp immobilized transcription
templates was achieved by ligation of 145 base pair free or nucleosomal DNA containing a
single engineered cisplatin intrastrand cross-link on either the template or coding strand with a
50 bp biotinylated promoter fragment with 9 nt overhangs. Analysis of resulting RNA transcript
length revealed that the T7 RNAP elongation complex can overcome the energy barrier to
nucleosome sliding caused by platinum intrastrand cross-links, but stalls when it reaches a PtDNA adduct placed on the DNA template strand. These results provide further evidence that
intrastrand cross-links of cisplatin inhibit transcription by creating a physical barrier that the
polymerase cannot pass.
192
References
(1)
Kirov, N.; Tsaneva, I.; Einbinder, E.; Tsanev, R. EMBO J 1992, 11, 1941-1947.
(2)
Studitsky, V. M.; Clark, D. J.; Felsenfeld, G. Cell 1994, 28, 371-382.
(3)
Studitsky, V. M.; Clark, D. J.; Felsenfeld, G. Cell 1995, 83, 19-27.
(4)
Protacio, R. U.; Widom, J. J Mol. Biol. 1996, 256, 458-472.
(5)
Bednar, J.; Studitsky, V. M.; Grigoryev, S. A.; Felsenfeld, G. Mol. Cell 1999, 4, 377-386.
(6)
Studitsky, V. M.; Kassavetis, G. A.; Geiduschek, E. P.; Felsenfeld, G. Science 1997, 278,
1960-1963.
(7)
Felsenfeld, G.; Clark, D.; Studitsky, V. M. Biophys. Chem. 2000, 86, 231-237.
(8)
Gottesfeld, J. M.; Belitsky, J. M.; Melander, C.; Dervan, P. B.; Luger, K. J. Mol. Biol.
2002, 321, 249-263.
(9)
Mohammad-Rafiee, F.; Kulic, I. M.; Schiessel, H. J. Mol. Biol. 2004, 344, 47-58.
(10)
Kireeva, M. L.; Walter, W.; Tschemajenko, V.; Bondarenko, V. A.; Kashlev, M.;
Studitsky, V. M. Mol Cell 2002, 9, 541-552.
(11)
Kireeva, M. L.; Hancock, B.; Cremona, G. H.; Walter, W.; Studitsky, V. M.; Kashlev, M.
Molecular Cell 2005, 18, 97-108.
(12)
Bondarenko, V. A.; Steele, L. M.; Ujvari, A.; Gaykalova, D. A.; Kulaeva, 0. I.;
Polikanov, Y. S.; Luse, D. S.; Studitsky, V. M. Mol. Cell 2006, 24, 468-479.
(13)
Orphanides, G.; LeRoy, G.; Chang, C. H.; Luse, D. S.; Reinberg, D. Cell 1998, 92, 105106.
(14)
Belotserkovskaya, R.; Oh, S.; Bondarenko, V. A.; Studitsky, V. M.; Orphanides, G.;
Reinberg, D. Science 2003, 301.
(15)
Belotserkovskaya, R.; Saunders, A.; Lis, J. T.; Reinberg, D. Biochim. Biophys. Acta
2004, 1677, 87-99.
(16)
Todd, R. C.; Lippard, S. J. Metallomics 2009, 1, 280-291.
(17)
Danford, A. J.; Wang, D.; Wang,
U.S.A. 2005, 102, 12311-12316.
(18)
Ober, M.; Lippard, S. J. J Am. Chem. Soc. 2007, 129, 6278-6286.
(19)
Ober, M.; Lippard, S. J. J Am. Chem. Soc. 2008, 130, 2851-2861.
Q.; Tullius,
193
T. D.; Lippard, S. J. Proc. Natl. Acad. Sci.
(20)
Wu, B.; Davey, C. A. Chem. Biol. 2008, 15, 1023-1028.
(21)
Luger, K. Chromosome Res. 2006, 14, 5-16.
(22)
Dhara, S. C. Indian J. Chem. 1970, 8, 193-194.
(23)
Todd, R. C.; Lovejoy, K. S.; Lippard, S. J. J. Am. Chem. Soc. 2007, 129, 6370-6371.
(24)
Dyer, P. N.; Edayathumangalam, R. S.; White, C. L.; Bao, Y.; Chakravarthy, S.;
Muthurajan, U. M.; Luger, K. Methods Enzymol. 2004, 375, 23-44.
(25)
Godde, J. S.; Wolffe, A. P. J Biol. Chem. 1995, 270, 27399-27402.
(26)
Jung, Y.; Lippard, S. J. J Biol. Chem. 2003, 278, 52084-52092.
(27)
Walter, W.; Studitsky, V. M. Methods 2004, 33, 18-24.
(28)
Wu, B.; Davey, C. A. Chem. Biol. 2008, 15, 1023-1028.
(29)
Gelasco, A.; Lippard, S. J. Biochemistry 1998, 37, 9230-9239.
(30)
Teuben, J.-M.; Bauer, C.; Wang, A. H.-J.; Reedijk, J. Biochemistry 1999, 38, 1230512312.
(31)
He, B.; Kukarin, A.; Temiakov, D.; Chin-Bow, S. T.; Lyakhov, D. L.; Rong, M.; Durbin,
R. K.; McAllister, W. T. J Biol. Chem. 1998, 273, 18802-18811.
(32)
Milligan, J. F.; Groebe, D. R.; Witherell, G. W.; Uhlenbeck, 0. C. Nucleic Acids Res.
1987, 15, 8783-88798.
(33)
Groebe, D. R.; Uhlenbeck, 0. C. Nucleic Acids Res. 1988, 16, 11725-11735.
(34)
Macdonald, L. E.; Zhou, Y.; McAllister, W. T. J.Mol. Biol. 1993, 232, 1030-1047.
(35)
Konarska, M. M.; Sharp, P. A. Cell 1989, 57, 423-43 1.
(36)
Nacheva, G. A.; Berzal-Herranz, A. Eur.J. Biochem. 2003, 270, 1458-1465.
(37)
Rong, M.; Durbin, R. K.; McAllister, W. T. J. Biol. Chem. 1998, 273, 10253-10260.
(38)
Schenbom, E. T.; Mierendorf Jr., R. C. Nucleic Acids Res. 1985, 13, 6223-6236.
194
RESEARCH SUMMARY AND PERSPECTIVES
Rational design of new platinum anticancer compounds with improved activity or novel
utility requires detailed knowledge of the mechanism of action of molecules in this family.
Research in this thesis is directed toward addressing two questions; (i) how adducts of platinum
complexes affect nucleic acid structure at the DNA and nucleosome level, and (ii) how these
structural deviations lead to inhibition of transcription, which correlates directly with tumor cell
death. We have advanced the field of research in two primary areas. First, the monofunctional
DNA adduct of pyriplatin, one member of a different class of active platinum compounds, was
structurally characterized. Combined with results from cell uptake, in vitro transcription, and
DNA repair assays, many details of the mechanism of action of active monofunctional platinum
complexes were revealed. Second, a more physiologically relevant model of cisplatin-DNA
damage was obtained, that of a 1,3-cis-{Pt(NH 3)2 }2+-d(GpTpG) intrastrand cross-link on DNA in
the nucleosome, via a 3.2-A resolution X-ray crystal structure. Nucleosome mobility and in vitro
transcription assays demonstrated that Pt-DNA adducts moderately inhibit histone translocation,
but that this effect does not play into the mechanism of transcription inhibition by cisplatin.
Future research on pyriplatin and other monofunctional platinum anticancer compounds
should focus on improving the activity of these complexes using the mechanistic insight
described here. The antineoplastic potential of pyriplatin has been demonstrated in animal tumor
models and in cell lines, but this molecule is much less potent than cisplatin. Utilizing orthosubstituted pyridine ligands could maximize steric interactions between the Pt-DNA adduct and
pol II bridge helix without altering the DNA-binding ability of the compound. Platinum(IV)
analogs of pyriplatin are currently being synthesized and investigated in our lab to improve cell
195
uptake, as well as to incorporate tumor-targeting and other anticancer agents. Use of nontraditional platinum compounds such as monofunctional, polyplatinum, or platinum(IV) agents
may be necessary in order to achieve a unique spectrum of activity against cancer.
The structural and functional studies of cisplatin-DNA cross-links on the nucleosome
reveal important information regarding the effects of platinum damage on higher order DNA
structure, as well as insight into the mechanism of transcription inhibition in vivo. The results
reported here provide strong evidence that disruption of chromatin dynamics can be overcome by
the cell, and that these effects do not contribute to blocking RNA synthesis in cancer cells. From
these data it can be inferred that ATP-dependent chromatin remodelers and RNA polymerase II
can act on platinated nucleosomes, but direct evidence has not yet been attained. Further
investigations in this area could focus on these eukaryotic enzymes.
196
Appendix A. Towards Separation of PtBP6-DNA Orientational Isomers by HPLC
197
Introduction
Studying the interactions between Pt-DNA adducts and other biological molecules is
important for understanding the mechanisms by which these DNA damaging agents disrupt
cellular processes.1 Investigations of protein selective recognition of Pt lesions are chief among
these concerns. Cisplatin intrastrand cross-links bend and unwind DNA in a manner similar to
that which occurs after protein binding,2 3 and a number of proteins, summarized in Chapter 1,
are known to recognize Pt-DNA adducts with specificity over naked DNA. Previous research has
focused on identifying these proteins in an effort to elucidate the mechanism of cellular response.
A cisplatin analog with a tethered benzophenone moiety (cis-[Pt(NH 3)(BP6)CI], PtBP6. see Fig.
A.J) was synthesized and attached to oligonucleotide probes. Benzophenone is a photoreactive
compound that forms cross-links through a radical mechanism.4 DNA probes were incubated
with cellular extracts, allowing nuclear proteins to interact with the platinum lesion.
The
samples were then irradiated with near UV (360 nm) light, forming a cross-link between the
tethered benzophenone and nearby proteins. Protein-DNA adducts were isolated by gel
electrophoresis and identified by mass spectrometry.
Using a short double stranded 25mer-
PtBP6 probe, several nuclear proteins were identified, including HMGB 1 and PARP- 1.5,6
HMGB 1 binds platinum-DNA lesions,3 and PARP- 1 is established as a DNA damage protein.7
The exact role of these proteins in cellular processing of cisplatin-DNA adducts has been
discussed but is not yet clear. 8
New probes were subsequently prepared that remove blunt ends, which are also targets
for proteins that respond to double-strand breaks, from the vicinity of the benzophenone site.
These probes include a 90mer dumbbell probe that contains no blunt ends, and a 157mer probe
198
that distances the blunt ends from the platinum site. Similar DNA-binding proteins were captured
using these complexes.9
CI
NW
C1
'YO
Pt
Pt
5'
5'
3'
3'
Figure A.1. (top) The structure of PtBP6. (bottom) depiction of the two orientational isomers of
PtBP6 on 14mer single-stranded DNA, where the benzophenone moiety is directed towards
either the 5' or 3' end of the oligonucleotide.
PtBP6 is capable of binding single-stranded DNA in two possible orientations in which
the benzophenone moiety is positioned towards either the 5' or 3' end of the oligonucleotide
strand (shown in Fig. A.1). Purification of these orientational isomers would provide a new tool
for studying protein binding of Pt-DNA adducts because one could obtain more structural
information about binding interactions if the photoactive probe location were precisely known.
This report describes attempts to resolve the orientational isomers of PtBP6 bound to 14mer
single-stranded DNA by reverse-phase HPLC. This project was halted before full separation of
the two products was achieved, but the progress is summarized here for to aid future work on
PtBP6-DNA probes that require isomerically pure material.
Experimental
Materials. PtBP6 was obtained from Dr. Datong Song, and its purity was confirmed by 1H NMR
spectroscopy (data not shown). Phosphoramidites and other reagents for DNA synthesis were
199
obtained from Glen Research. All other reagents and solvents were purchased from commercial
sources and used without further purification. DNA syntheses were performed on an Applied
Biosystems Model 392 DNA/RNA synthesizer at a 1.0 gmol scale. HPLC applications were
performed using a Waters 600 system. Atomic absorption spectroscopy was performed on a
Perkin-Elmer AAnalyst 300 system.
1H
NMR spectra were obtained on a Bruker 300 MHz
NMR spectrometer. UV-VIS spectra were obtained on a HP 8453 UV-visible spectrometer.
Platination of 14mer with PtBP6. The 14mer oligonucleotide 5'-TTCACCGTGATTCC-3' was
synthesized and purified as described in Chapter 5 of this thesis. Pt(BP6)(NH 3)ClI (0.98 mg,
1.37 pmol dissolved in 90 pL DMF) was allowed to react with AgNO 3 (1.95 equiv dissolved in
10 pL ddH 2O) for 4 h at ambient temperature in the dark. After AgCl precipitate was removed
by centrifugation,
50 nmol of 14mer were combined with 2 equiv of activated PtBP6
supernatant in 500 pL of 10 mM sodium phosphate pH 6.8, and the solution was allowed to react
at 37 'C overnight.
The products were analyzed and purified by HPLC using the methods
described below. Peaks were collected manually, then were dialyzed against 3 changes of 2 L of
water, lyophilized to dryness, re-dissolved in 1 mL ddH2 0, and analyzed by UV-VIS and AA
spectroscopy. Peaks with a Pt/DNA ratio of -1.0 were analyzed by Maxam-Gilbert sequencing
to confirm platination at the two guanine sites; binding at the N7 position blocks cleavage of the
Maxam-Gilbert G-reaction (data not shown).
Resolution of orientational isomers by HPLC. Initial attempts to resolve PtBP6-14mer
orientational isomers by RP-HPLC were conducted using a Vydac Protein/Peptide C4 column
and a gradient of 100 mM triethylammonium acetate (TEAA) pH 7 and acetonitrile. Method
200
details and a sample chromatogram are shown in Fig. A.2. After insufficient resolution was
obtained with this column, the more hydrophobic Vydac Protein/Peptide C18 column was used.
Two unresolved peaks were observed, which became better separated when the concentration of
TEAA was increased from 50 mM to 100 mM. Details of this method, along with chromatograms, are given in Figs. A.3 and A.4.
Method Details
Column:
Column Temp:
Flow rate:
Detection:
Mobile phase A:
Mobile phase B:
Vydac Protein/Peptide C4, 4 x 250 mm, 5 pm
ambient
1.0 mL/min
UV @ 260 nm
50 mM triethylammonium acetate pH 7.0
acetonitrile
Gradient
Time: 0
40
45
45.1
50
%B:
60
60
5
5
5
14mer
0.20-
0.15PtBP6-14mer
0.10
0.00
5.00
10.00
15.00
Minutes
20.00
25.00
Figure A.2. HPLC method and chromatogram for purification of PtBP6-14mer from
unplatinated oligonucleotide using a C4 reverse-phase column. No resolution of PtBP6-14mer
orientational isomers is observed.
201
Method Details
Column:
Column Temp:
Flow rate:
Detection:
Mobile phase A
Mobile phase B
Vydac Protein/Peptide C18, 4.6 x 250 mm, 5 pm
ambient
1.0 mL/min
UV @260 nm
50 mM triethylammonium acetate pH 7.0
acetonitrile
Gradient
Time: 0
%B: 1(
25.1
80
2.00
4.00
30
80
30.1
10
6.00
8.00
35
10
10.00
12.00
14.00
Minutes
Figure A.3. HPLC method and chromatogram for purification of PtBP6-14mer from
unplatinated oligonucleotide using a C18 reverse-phase column and 50 mM TEAA. Minimal
resolution of PtBP6-14mer orientational isomers is observed.
202
Method Details
Column:
Column Temp:
Flow rate:
Detection:
Mobile phase A
Mobile phase E
Vydac Protein/Peptide C18, 4.6 x 250 mm, 5 pm
ambient
1.0 mL/min
UV @ 260 nm
7.5% acetonitrile/100 mM TEAA pH 7.0
50% acetonitrile/100 mM TEAA pH 7.0
Gradient
Time: 0
%B: 0
31
100
36
100
37
0
0.W
5.00
10.00
15.00
20.00
25.00
Minutes
30.00
35.00
40.00
Figure A.4. HPLC method and chromatogram for purification of PtBP6-14mer from
unplatinated oligonucleotide using a C18 reverse-phase column and 100 mM TEAA. Decent
resolution of PtBP6-14mer orientational isomers is observed.
Results and Discussion
Reverse-phase HPLC was used to purify PtBP6-14mer from unplatinated DNA and to
attempt to resolve the two orientational isomers of this compound. The benzophenone moiety
adds sufficient hydrophobicity to the oligonucleotide so that separation of platinated and
unplatinated DNA is simple. Triethylammonium acetate is used as a mobile phase additive; the
cations associate with the negatively charged phosphodiester backbone, and the alkylated
ammonium ions further increase the DNA hydrophobicity. However, differences in
hydrophobicity between the two orientational isomers of PtBP6-14mer are much more subtle,
203
and conformational differences must be relied on to separate the two species, since the chemical
compositions are identical.
Resolution of the two isomers was not possible on a C4 column; the platinated DNA
eluted as a single peak (see Fig. A.2). Using the more hydrophobic C18 column, the desired peak
split into two approximately equal peaks, but they were not fully resolved. Both peaks contained
one Pt per DNA strand. By increasing the TEAA concentration in the aqueous mobile phase,
better (but still not baseline) -resolution was obtained. These two results collectively suggest that
by increasing the hydrophobic adsorption interactions between PtBP6-14mer molecules and the
solid phase, the Pt-DNA complex can be separated into two distinct species. However, these
species have not yet been confirmed as the two orientational isomers. NMR spectroscopic studies
were initiated to identify the two species.9
Conclusions
The photoreactive Pt-DNA species PtBP6-14mer was synthesized, and attempts to isolate
and purify the two orientational isomers of this compound were undertaken. Advances were
made utilizing reverse-phase HPLC and a C18 column, but the project was halted before
complete characterization of the products was performed. These data could be useful for future
projects using this Pt-DNA probe. Using a specific orientational isomer could reveal more
structural information about the nature of protein recognition of Pt-DNA adducts.
204
References
(1)
Wang, D.; Lippard, S. J. Nat. Rev. Drug Discov. 2005, 4, 307-320.
(2)
Coin, F.; Frit, P.; Viollet, B.; Salles, B.; Egly, J.-M. Mol. Cell BIol. 1998, 18, 3907-3914.
(3)
Ohndorf, U.-M.; Rould, M. A.; He,
712.
(4)
Dorman, G.; Prestwich, G. D. Biochemistry 1994, 33, 5661-5673.
(5)
Zhang, C. X.; Chang, P. V.; Lippard, S. J. J. Am. Chem. Soc. 2004, 126, 6536-6537.
(6)
Guggenheim, E. R.; Xu, D.; Zhang, C. X.; Chang, P. V.; Lippard, S. J. Chem. Biochem.
2009, 10, 141-157.
(7)
Schreiber, V.; Dantzer, F.; Ame, J.-C.; de Murcia, G. Nat. Rev. Mol. Cell Biol. 2006, 7,
517-528.
(8)
Todd, R. C.; Lippard, S. J. Metallomics 2009, 1, 280-291.
(9)
Guggenheim, E. R., Massachusetts Institute of Technology, 2008.
Q.; Pabo, C. 0.; Lippard, S. J. Nature 1999, 399, 708-
205
Appendix B. Crystallization Attempts of a DNA 11mer Duplex Containing a Site-Specific
1,3-cis-{Pt(NH 3)2 }2 -d(GpTpG) Lesion
206
Introduction
A summary of the structural studies of double-stranded DNA containing cisplatin damage
sites is provided in Chapter 1. All of the major structural adducts of cis-diamminedichloroplatinum(II) have been characterized
either by X-ray crystallography, 1-3 NMR
spectroscopy, 4 -9 or by both methods. Noticeably absent from this list is the solid-state structure of
the 1,3-cis-{Pt(NH 3) 2}2+-d(GpTpG) intrastrand cross-link on DNA. Although NMR solution
structures of this adduct on duplex DNA have been solved (see Fig. B.1),
47
no X-ray crystal
structure of what might be the major adduct of carboplatin10 exists. In the NMR structure the
duplex is bent by ~30' and the double helix displays local unwinding and widening of the minor
groove, similarly to features of the structure of the 1,2-d(GpG) cross-link. The 1,3-d(GpTpG)
adduct differs, however, in that base pairing of the 5' G*-C, where the asterisk denotes a
platination site, is disrupted and the internal thymidine of the adduct is extruded from the minor
groove.
Figure B.1. Stereo view of the NMR solution structure of duplex DNA containing a cisplatin
1,3-cis-{Pt(NH 3)2}2+-d(GpTpG) intrastrand cross-link. The platinum moiety is bound at the N7
positions of each guanine, with the internal thymine pointed outside the double helix.
207
Differences in the structures of the 1,2-d(GpG) cross-link solved by these two
28
methodologies underscore the importance of analyzing these complexes by both techniques. 'ii
The X-ray crystal structure of the 1,3-(GpTpG) adduct on the nucleosome is described in
Chapter 5, but a crystallographic model of this platinum damage site on naked DNA is still
desired for comparison to both the NMR solution structure and the nucleosome model. Thus,
attempts to crystallize the 1,3-cis-{Pt(NH 3)2}"2 -d(GpTpG) intrastrand cross-link on an 1lmer
DNA duplex were undertaken. The DNA sequence, 5'-CTCTG*TG*TCTC-3', is identical to
that used in the NMR solution structure determination. Highly pure material was isolated and
crystallization studies were initiated, but diffraction-quality crystals were never obtained. The
results described here will be useful to someone wishing to continue this project and complete a
key missing piece of the cisplatin-DNA adduct structure profile.
Experimental
Materials. Phosphoramidites, columns, and other reagents for solid-phase oligonucleotide
synthesis were purchased from Glen Research. Potassium tetrachloroplatinate(II), which was
2
used to synthesize cisplatin according to published procedures,' was a gift from Engelhard
(Iselin, NJ, now BASF). All other reagents were purchased from commercial suppliers and used
without further purification. Oligonucleotides were prepared in-house using an Applied
Biosystems Model 392 DNA/RNA Synthesizer. Liquid chromatography was performed with an
Agilent 1200 series HPLC equipped with a temperature-controlled autosampler and automated
fraction collector. UV-Vis spectra were collected on a Hewlett-Packard 8453 spectrophotometer.
208
Synthesis and purification of 11mer duplex containing the 1,3-cis-{Pt(NH 3)2 }2 -d(GpTpG)
cross-link. The oligonucleotides 5'-d(CTCTGTGTCTC)-3' (11-ts) and its complementary
strand (11-bs) were chemically synthesized by standard solid-phase methods
3
on a 10 pImol
scale with trityl groups intact. The strands were purified by reversed-phase HPLC on an Agilent
SB-300, 9.4 x 250 mm column using method RCT.005S (see Appendix C). Trityl groups were
removed in 80% acetic acid for 30 min at room temperature, and the oligonucleotides were
precipitated with isopropanol and desalted with Sep-Pak C18 cartridges. Yields were determined
by UV-Vis spectroscopy to be 1.27 pmol (13%) and 0.83 pmol (8%) for 11-ts and 11-bs,
respectively. Platination reactions were carried out as described in Chapter 4, yielding 434 nmol
of 11-Pt (34%). The site-specifically platinated duplex was prepared by combining equimolar
amounts of 11-Pt and 11-bs in 200 pL of 200 mM LiCl, 100 mM HEPES pH 7.0, and 50 mM
MgCl 2 , heating to 70 'C for 10 min, and cooling to 4 'C over 2.5 h. The solution was purified by
ion exchange HPLC on a Dionex DNA-Pac PA-100 9 x 250 mm column by method RCT.003S,
and desalted by C18 SPE. Chromatograms of final purification steps are shown in Fig. B.2.
Crystallization attempts of Pt-11mer duplex. Crystallization trays were set up using the
hanging drop vapor diffusion method. Starting conditions were obtained from a survey of
crystallization conditions for other Pt-DNA duplexes. 3 ,14-1 6 In one set of conditions, 4 ptL
hanging drops initially containing 0.2 mM Pt-DNA, 50 mM sodium cacodylate pH 6.5, 2 mM
spermine, 10-60 mM MgCl 2 , and 5-30% (v/v) 2-methyl-2,4-pentanediol (MPD) were allowed to
equilibrate against a reservoir of 30% MPD. In another tray, crystallization solutions contained
50 mM sodium cacodylate pH 6.5, 2 mM spermine, 80-130 mM Mg(OAc) 2, and 10-35% (w/v)
polyethylene glycol (PEG) 4000. Hanging drops contained 2 pL of 0.4 mM DNA in water and 2
209
pL of crystallization solution. All solutions were prepared and sterile filtered immediately prior
to use. Crystallization trays were stored at 4 'C in the dark.
2500
2000
4 1500
1
1000
500
25
20
15
10
5
Time (min)
2500
2000
1500
4iC~
1000
)0
0
0
5
10
15
20
Time (min)
25
30
35
40
Figure B.2. (top) Purification of 1 imer platination reaction by ion-exchange HPLC. Peak 1
represents platinated DNA, and peak 2 the unplatinated starting material. (bottom) Purification of
11-Pt DNA duplex (peak 3) after annealing of the platinated oligonucleotide with its
complementary strand.
210
Results and Discussion
Synthesis/purification of 11mer duplex w/ 1,3-cis-{Pt(NH 3)2}2 -d(GpTpG) adduct. The
oligonucleotides 11-ts and 11-bs were obtained in decent yield after HPLC purification. The
platination reaction proceeded cleanly (Fig. B.2, top), aided in part by design of the DNA strand
to contain exactly two guanine reactive sites and no other purine bases. Annealing the platinated
strand and its complement to form the 1lImer double-stranded DNA was initially problematic
due to its low melting temperature, calculated to be 34 'C without the Pt cross-link at 50 mM
NaCl, 17 but the duplex was eventually obtained by increasing the LiCl concentration to 200 mM
and the oligonucleotide concentration to -2 mM. Double-stranded DNA structure was confirmed
by the later retention time of the product peak on the HPLC chromatogram (Fig. B.2, bottom).
However collection of the product after HPLC purification results in significant dilution of the
sample, which may have resulted in dissociation of the duplex. Two possible ways to address
this problem include annealing the 11 mer duplex and crystallizing without HPLC purification
and increasing the DNA strand length to 12 or 13 base pairs to raise the melting temperature. The
former solution would remove the possibility that dilution of the DNA sample would cause loss
of the double-stranded structure, but may introduce single-stranded impurities into the sample
that could prohibit crystallization. The latter solution would most likely ensure stability of the
double-stranded DNA by increasing Td, but would introduce its own possible drawbacks. A
dodecamer duplex containing the 1,3-GpTpG adduct would be asymmetric, unlike the 1,2intrrastrand cross-link, which has been studied on 12mer DNA, and could complicate
crystallization. The 13mer DNA duplex would be symmetric, but increasing the DNA length
increases the flexibility of the molecule, which introduces different crystallization challenges.
211
Crystallization attempts of Pt-11mer duplex. Several of the explored conditions containing
~14% MPD produced microcrystalline material of the Pt-DNA complex, but no diffractionquality single crystals were ever obtained. This study utilized crystallization conditions used to
solve high-resolution X-ray structures of platinum-DNA duplexes. Subsequent efforts should
focus on broadening the crystallization condition matrix, perhaps using the Hampton nucleic acid
screen Natrix," to find the appropriate conditions for obtaining diffraction-quality crystals. The
extruded internal thymine base in the platinum cross-link may sufficiently alter crystal-packing
interactions so that crystallization under the more conventional conditions was not possible.
Conclusion
Double-stranded 11Imer DNA containing the 1,3-cis-{Pt(NH3)2} 2 -d(GpTpG) intrastrand
cross-link of cisplatin was synthesized and purified in high quantities. Crystallization studies
were initiated but diffraction-quality crystals were never obtained. Two possible reasons are that
single-stranded DNA was present in the sample, or that the structure of the Pt-DNA duplex was
sufficiently different that crystallization under previously successful conditions was not favored.
Possible solutions for removing single-stranded contaminants or promoting crystal formation are
presented here.
212
References
(1)
Takahara, P. M.; Rosenzweig, A. C.; Frederick, C. A.; Lippard, S. J. Nature 1995, 377,
649-652.
(2)
Takahara, P. M.; Frederick, C. A.; Lippard, S. J. J Am. Chem. Soc. 1996, 118, 1230912321.
(3)
Coste, F.; Malinge, J.-M.; Serre, L.; Shepard, W.; Roth, M.; Leng, M.; Zelwer, C. Nucleic
Acids Res. 1999, 27, 1837-1846.
(4)
Van Garderen, C. J.; Van Houte, L. P. A. Eur. J Biochem. 1994, 225, 1169-1179.
(5)
Yang, D.; van Boom, S. S. G. E.; Reedijk, J.; van Boom, J. H.; Wang, A. H.-J.
Biochemistry 1995, 34, 12912-12920.
(6)
Huang, H.; Zhu, L.; Reid, B. R.; Drobny, G. P.; Hopkins, P. B. Science 1995, 270, 18421845.
(7)
Teuben, J.-M.; Bauer, C.; Wang, A. H.-J.; Reedijk, J. Biochemistry 1999, 38, 1230512312.
(8)
Marzilli, L. G.; Saad, J. S.; Kuklenyik, Z.; Keating, K. A.; Xu, Y. J. Am. Chem. Soc.
2001, 123, 2764-2770.
(9)
Wu, Y.; Bhattacharyya, D.; King, C. L.; Baskerville-Abraham, I.; Huh, S.-H.; Boysen,
G.; Swenberg, J. A.; Temple, B.; Campbell, S. L.; Chaney, S. G. Biochemistry 2007, 46,
6477-6487.
(10)
Blommaert, F. A.; van Dijk-Knijnenburg, H. C. M.; Dijt, F. J.; den Engelse, L.; Baan, R.
A.; Berends, F.; Fichtinger-Schepman, A. M. J. Biochemistry 1995, 34, 8474-8480.
(11)
Todd, R. C.; Lippard, S. J. J Inorg. Biochem. 2010, 104, in press.
(12)
Dhara, S. C. Indian J. Chem. 1970, 8, 193-194.
(13)
Caruthers, M. H. Acc. Chem. Res. 1991, 24, 278-284.
(14)
Silverman, A. P.; Bu, W.; Cohen, S. M.; Lippard, S. J. J Biol. Chem. 2002, 277, 4974349749.
(15)
Komeda, S.; Moulaei, T.; Woods, K.; Chikuma, M.; Farrell, N.; Williams, L. J Am.
Chem. Soc. 2006, 128, 16092-16103.
(16)
Lovejoy, K. S.; Todd, R. C.; Zhang, S.; McCormick, M. S.; D'Aquino, J. A.; Reardon, J.
T.; Sancar, A.; Giacomini, K. M.; Lippard, S. J. Proc.Natl. Acad Sci. U.S.A. 2008, 105,
8902-8907.
(17)
Kibbe, W. A. Nucleic Acids Res. 2007, 35, W43-W46.
213
(18)
Scott, W. G.; Finch, J. T.; Grenfell, R.; Fogg, J.; Smith, T.; Gait, M. J.; Klug, A. J. Mol.
Biol. 1995, 250, 327-332.
214
Appendix C: HPLC Methods for Purification and Analysis of Platinated Oligonucleotides
Legend
P = preparative HPLC method
S = semi-preparative HPLC method
A = analytical HPLC method
M
=
HPLC/mass spectrometry method
215
Method #:
RCT.001P
Description:
Purification of dimethoxytrityl-containing short oligonucleotides
from un-tritylated failure sequences by preparative reverse-phase HPLC
Method Details
Column:
Column Temp:
Flow rate:
Detection:
Mobile phase A:
Mobile phase B:
Gradient
Time: 0
%B: 20
Vydac C18, 16 x 250 mm, 5 pm
ambient
10.0 mL/min
UV @ 260 nm
50 mM triethylammonium acetate pH 7.0
acetonitrile
11.1
20
15
20
Approximate retention times (boldface indicates desired product)*
1) Failure sequences:
2) DMT-oligos (12-14 nt):
< 5 min
6-8 min
*Actual retention time can vary based on oligonucleotide length and column condition.
216
Method #:
RCT.002S
Description:
Purification of 12mer DNA platination reactions by semi-preparative ionexchange HPLC
Method Details
Column:
Column Temp:
Flow rate:
Detection:
Mobile phase A
Mobile phase E
Dionex DNA-Pac PA-100, 9 x 250 mm
ambient
4.0 mL/min
UV @ 260 nm
90:10 (25 mM NH 40Ac pH 5.2):acetonitrile + 0.2 M NaCl
90:10 (25 mM NH 40Ac pH 5.2):acetonitrile + 0.4 M NaCl
Gradient
Time: 0
%B: 0
36
0
35
100
Approximate retention times (boldface indicates desired product)*
<11 min
13-14 min
24 min
Multiple-Pt-containing species:
Pt-12mer:
12mer starting material:
*Actual retention time can vary based on column condition.
Sample Chromatogram
1800 1600 1400 1200 1000 C
800 600 400 200
0
0
5
10
20
15
Time (min)
217
25
30
35
40
Method #:
RCT.003S
Description:
Purification of dodecamer duplex DNA containing a single platinum-DNA adduct
by semi-preparative ion-exchange HPLC
Method Details
Column:
Column Temp:
Flow rate:
Detection:
Mobile phase A
Mobile phase B
Gradient
Time: 0
%B: 0
Dionex DNA-Pac PA-100, 9 x 250 mm
ambient
4.0 mL/min
UV @ 260 nm
90:10 (25 mM NH4 0Ac pH 5.2):acetonitrile + 0.2 M LiCi
90:10 (25 mM NH4 0Ac pH 5.2):acetonitrile + 0.4 M LiCi
35
100
36
0
Approximate retention times (boldface indicates desired product)*
1) Single-stranded DNA:
2) Duplex Pt-DNA:
18-25 min
32 min
*Actual retention time can vary based on column condition.
Sample Chromatogram
218
-11 __
4
..........
Method #:
RCT.004A
Description:
Analysis and purification of 14mer DNA platination reactions with cisplatin by
ion-exchange HPLC
Method Details
Column:
Column Temp:
Flow rate:
Detection:
Mobile phase A:
Mobile phase B:
Gradient
Time: 0
%B: 5
Agilent Zorbax Oligo, 6.2 x 80 mm
45 *C
1.0 mL/min
UV @ 260 nm
80:20 (20 mM sodium phosphate pH 7.0):acetonitrile
A + 1.0 M NaCl
70
25
75
80
70.1
80
75.1
5
80
5
Approximate retention times (boldface indicates desired product)*
1) Pt-14mer
2) 14mer
26-29 min
46 min
*Actual retention time can vary based on column condition.
Sample Chromatogram
120
100
~80
60
40
20
0~
0
10
20
40
30
Time (min)
219
50
60
70
80
Method #:
RCT.005S
Description:
Purification of dimethoxytrityl-containing short oligonucleotides
from un-tritylated failure sequences by semi-preparative reverse-phase HPLC
Method Details
Column:
Vydac Cl8, 9 x 250 mm, 5 pm or Agilent C18 SB-300, 9.4 x 250
mm, 5 pm
ambient
4.0 mL/min
UV @ 260 nm
50 mM triethylammonium acetate pH 7.0
acetonitrile
Column Temp:
Flow rate:
Detection:
Mobile phase A
Mobile phase B
Gradient
Time: 0
%B: 20
11.1
20
15
20
Approximate retention times (boldface indicates desired product)*
1) Failure sequences:
2) DMT-oligos (12-14 nt):
< 5 min
6-8 min
*Actual retention time can vary based on oligonucleotide length and column condition.
Sample Chromatogram
3500 ,
3000 -
2500
W2000UU
1500
1000 -
500
0
0
-
I1
0
2
[,
4
-
-
6
8
Time (min)
220
10
12
14
Method #:
RCT.006S
Description:
Purification of 14mer DNA platination reactions by semi-preparative ionexchange HPLC
Method Details
Column:
Column Temp:
Flow rate:
Detection:
Mobile phase A:
Mobile phase B:
Gradient
Time:30
20
%B:
Dionex DNA-Pac PA-100, 9 x 250 mm
35 0 C
4.0 mE/mm
UV @ 260 nm
90:10 (25 mM sodium acetate pH 5.2):acetonitrile
A + 0.5 M NaCl
39
100
38
55
5
40
49
20
44
20
43
100
Approximate retention times (boldface indicates desired product)*
<12 min
16-17 min
37 min
1) Multiple-Pt-containing species:
2) Pt-14mer:
3) 14mer starting material:
*Actual retention time can vary based on column condition.
Sample Chromatogram
300
250
200
E
150
A 100
50
j
L-
0
0
5
10
15
20
25
Time (min)
221
30
35
40
45
50
Method #:
RCT.007A
Description:
Purification of 14mer DNA platination reactions by analytical ionexchange HPLC
Method Details
Column:
Column Temp:
Flow rate:
Detection:
Mobile phase A:
Mobile phase B:
Gradient
Time: 0
%B: 5
Agilent Zorbax Oligo, 6.2 x 80 mm
45 0C
1.0 mL/min
UV @ 260 nm
80:20 (20 mM sodium phosphate pH 7.0):acetonitrile
A + 1.0 M NaCl
40
16.5
40.1
80
45
80
45.1
5
50
5
Approximate retention times (boldface indicates desired product)*
1) Multiple-Pt-containing species:
2) Pt-14mer:
3) 14mer starting material:
<8 min
10-12 min
18 min
*Actual retention time can vary based on column condition.
Sample Chromatogram
2500
2000
D
1500
1000
500
0
5
10
15
20
25
Time (min)
222
30
35
40
45
Method #:
RCT.008A
Description:
Analysis of platinated oligonucleotides digested with nuclease SI/P 1 and calf
intestinal phosphatase by reverse-phase HPLC
Method Details
Column:
Column Temp:
Flow rate:
Detection:
Mobile phase A
Mobile phase B
Supelcosil LC-18-S, 4.6 x 250 mm, 5 tm
ambient
1.0 mL/min
UV @ 260 nm
100 mM sodium acetate pH 5.2
methanol
Gradient
Time: 0
%B: 5
45.1
5
50
5
Note: Dependingon the Pt-DNA adduct, it may be necessary to modify the gradient.
Retention time*
7 min
15 min
17 min
27 min
1) deoxycytidine
2) deoxyguanosine
3) thymidine
4) deoxyadenosine
Relative response factor
1.0
1.6
1.2
2.0
*Actual retention time can vary based on column condition.
SamDle Chromatogram
2500
2000
1500
1000
500
0
10
30
20
Time (min)
223
40
-ZI.1-1--- 1-
Method #:
RCT.009M
Description:
Analysis of platinated oligonucleotides digested with nuclease SI/PI and calf
intestinal phosphatase by reverse-phase HPLC/ESI-MS
Method Details
Column:
Column Temp:
Flow rate:
Detection:
Mobile phase A
Mobile phase B
Supelcosil LC-18-S, 2.1 x 250 mm, 5 pm
ambient
0.2 mL/min
UV @ 260 nm, ESI-MS (-) mode
100 mM sodium acetate pH 5.2
methanol
Gradient
Time: 0
%B: 5
45.1
5
50
5
Note: Depending on the Pt-DNA adduct, it may be necessary to modify the gradient.
Relative response factor
1.0
1.6
1.2
2.0
Retention time*
7 min
15 min
17 min
27 min
deoxycytidine
deoxyguanosine
thymidine
deoxyadenosine
*Actual retention time can vary based on column condition.
Sample Chromatogram
2500
2000
1500
1000
500
0
10
20
30
Time (min)
224
40
Method #:
RCT.009M (cont.)
Description:
Analysis of platinated oligonucleotides digested with nuclease SI/P 1 and calf
intestinal phosphatase by reverse-phase HPLC/ESI-MS
Sample Mass Spectra
Inten S
x10 4
3
deoxycytidine
2284
287.7
389.9
1
0
200
Inten
x105
300
400
500
703
600
800
rnt
deoxyguanosine
2661
4
3
2
3479
0'
200
Inten
x10
6
5329
400
300
600
500
29thymidine
264,9
700
800
930
M&
90
m/I
5290
10
08
06
04
02
00
200
400
300
500
$00
Inten S I
x1C 5
700
800
deoxyadenosine
3102
4
3
2
1
800
400
225
m
Method #:
RCT.010M
Description:
Electrospray mass spectrometry of oligonucleotides with on-line HPLC desalting
Method Details
Column:
Column Temp:
Flow rate:
Detection:
Mobile phase A
Mobile phase B
:
:
Gradient
Time: 0
%B: 5
30
50
Agilent Extend C18, 2.1 x 150 mm, 3.5 pm
ambient
0.2 mL/min
ESI-MS (-) mode
5 mM NH 40Ac pH 5.2
acetonitrile
30.1
80
35
80
35.1
5
40
5
SamDle Mass SDectrum
hot~ns'.
x 104
-4
C aIc: 4425.8 Da
Obs: 4424.9 ± 1.7 D a
.7
I[
-7
200
4W
OW
800
226
1000
BIOGRAPHY
Ryan Christopher Todd was born on August 30, 1981 to Elaine S. O'Donnell and W.
Alan Todd in Abington, PA. He attended Perkiomen Valley High School (Collegeville, PA) and
graduated from Johns Hopkins University in 2003 with a B.A. in chemistry, during which time
he studied oxygen atom transfer reactivity of mangenese(III) and iron(III) triazacorrole
complexes under the supervision of Professor David P. Goldberg. He worked for two years as an
analytical chemist at Merck & Co., Inc. before beginning studies at MIT in 2005 in the
laboratory of Professor Stephen J. Lippard. As a graduate student he recieved the Koch Institute
Graduate Fellowship, David A. Johnson Graduate Student Summer Fellowship, and Strem
Graduate Student Summer Fellowship.
Ryan met his wife Moira in 2002 when they were classmates at Johns Hopkins and
became next-door neighbors. They were married in July 2007, and currently live in Somerville,
MA.
227
RYAN C. TODD
EDUCATION
2005 - 2010
Massachusetts Institute of Technology, Cambridge, MA
Ph.D. in Biochemistry
Research Advisor: Professor Stephen J. Lippard
1999 - 2003
Johns Hopkins University, Baltimore, MD
B.A. in Chemistry
Undergraduate Research Advisor: Professor David P. Goldberg
RESEARCH EXPERIENCE
2010
Research Assistant - Massachusetts Institute of Technology, Cambridge, MA
- Used X-ray crystallography, mass spectrometry, HPLC, gel electrophoresis, and
other techniques to study structural and functional consequences of binding of
platinum anticancer compounds to free and nucleosomal DNA.
2001 - 2003
Undergraduate Research Assistant - Johns Hopkins University, Baltimore, MD
- Synthesized, characterized, and investigated the oxygen atom transfer reactivity of
iron(III) and manganese(III) corrolazine complexes, and identified the oxidation
products of different substrates by gas chromatography.
2005
-
PROFESSIONAL EXPERIENCE
2005 - 2006
Teaching Assistant - Massachusetts Institute of Technology, Cambridge, MA
- Instructed an introductory chemistry laboratory course for non-chemistry majors.
- Provided assistance with experiments, wrote and graded quizzes, and tutored
students on relevant chemical principles.
2003 - 2005
Chemist - Merck & Co., Inc., West Point, PA
-
Developed, validated, and documented analytical HPLC methods for the assay of
drug entities and their potential degradates in research formulations to support
clinical release and stability testing for IND and NDA filings.
Provided analytical support for formulation and process development.
PUBLICATIONS
1)
Todd, RC, Lippard, SJ (2010) Structure of duplex DNA containing the cisplatin 1,2-{Pt(NH 3)2}2d(GpG) cross-link at 1.77 A resolution. J. Inorg. Biochem. in press.
2)
Todd, RC, Lippard, SJ (2009) Inhibition of Transcription by Platinum Antitumor Compounds.
Metallomics 1:280-291.
3)
Todd, RC, Lippard, SJ (2009) X-Ray Crystal Structure of a Monofunctional Platinum-DNA
Adduct, cis-{Pt(NH 3)2(pyridine)}2 * Bound to Deoxyguanosine in a Dodecamer Duplex. In: Bonetti,
A, Howell, SB, Leone, R, Muggia, F, eds. Platinum and Other Heavy Metal Compounds:
Molecular Mechanisms and Clinical Applications. Totowa: Humana Press; 67-72.
4)
Lovejoy, KS*, Todd, RC*, Zhang, S, McCormick, MS, D'Aquino, JA, Reardon, JT, Sancar, A,
Giacomini, KM, Lippard, SJ (2008) cis-Diammine(pyridine)chloroplatinum(II), a Monofunctional
Platinum(II) Antitumor Agent: Uptake, Structure, Function, and Prospects. Proc. Natl. Acad. Sci.
USA 105:8902-8907. (*Authors contributedequally to the paper)
5)
Todd, RC, Lovejoy, KS, Lippard, SJ (2007) Understanding the Effect of Carbonate Ion on
Cisplatin Binding to DNA. J. Am. Chem. Soc. 129:6370-6371.
228
6)
Wang, SH, Mandimutsira, BS, Todd, R, Ramdhanie, B, Fox, JP, Goldberg, DP (2004)
Catalytic Sulfoxidation and Epoxidation with a Mn(III) Triazacorrole: Evidence for A "Third
Oxidant" in High-Valent Porphyrinoid Oxidations. J. Am. Chem. Soc. 126:18-19.
7)
Mandimutsira, B, Ramdhanie, B, Wang, H, Todd, RC, Zareba, AA, Czernuszewicz, RS,
Goldberg, DP (2002) A Stable Manganese(V) Oxo-Corrolazine Complex. J. Am. Chem. Soc. 124:
15170-15171.
PRESENTATIONS AND POSTERS
1)
Structural and Functional Analysis of a Cytotoxic Monofunctional Platinum-DNA Adduct. MIT
Koch Institute Focus Seminar Series. Cambridge, MA (2007).
2)
X-Ray Crystal Structure of a Monofunctional Platinum-DNA Adduct, cis{Pt(NH 3)2(pyridine)} 2
Bound to Deoxyguanosine in a Dodecamer Duplex (poster). Xth International Symposium on
Platinum CoordinationCompounds in Cancer Chemotherapy.Verona, Italy (2007).
HONORS AND AWARDS
2009
2008
2008
2008
2007
2006
2003
2002
2002
2000
Strem Graduate Student Summer Fellowship
Koch Institute Graduate Fellowship
David A. Johnson Graduate Student Summer Fellowship
IPMI Gemini Graduate Student Award
Xth ISPCC: 1" Place Poster Prize
MIT Student Teaching Award
Phi Beta Kappa
Howard Hughes Summer Research Fellowship Program
Kilpatrick Prize for excellence in chemistry
Land Scholarship for excellence in chemistry
229