Analysis of the Structural Changes Caused by Positive DNA
Supercoiling
by-
Marita Christine Barth
B.S. Bioresource Research
Oregon State University, 1998
Submitted to the Division of Biological Engineering in partial fulfillment of the
requirements for the degree of
Doctor of Philosophy in Macromolecular Biochemistry and Biophysics
at the
MASSACHUSETTS INSTITUTE OF TECHNOLOGY
February 2007
C 2007 Massachusetts Institute of Technology. All rights reserved.
Signature of Author:
Division of Biological Engineering
January 12, 2007
Certified by:
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Peter C. Dedon
Professor of Toxicology and Biological Engineering
Thesis Supervisor
Accepted by:
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This doctoral thesis has been examined by a committee of the Division of Biological
Engineering as follows:
Associate Professor Bevin P. Engelward
Chairman
Professor Peter C. Dedon
Supervisor
Professor John M. Essigmann
Dr. Richard J. Roberts
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---
Analysis of the Structural Changes Caused by Positive DNA
Supercoiling
by
Marita Christine Barth
Submitted to the Division of Biological Engineering on January 12, 2007 in partial
fulfillment of the requirements for the degree of Doctor of Philosophy in Macromolecular
Biochemistry and Biophysics
ABSTRACT
The procession of helix-tracking enzymes along a DNA molecule results in the
formation of supercoils in the DNA, with positive supercoiling (overwinding) generated
ahead of the enzyme, and negative supercoiling (underwinding) in its wake. While the
structural and physiological consequences of negative supercoiling have been well
studied, technical challenges have prevented extensive examination of positively
supercoiled DNA. Studies suggest that at sufficiently high levels of overwinding, DNA
relieves strain by adopting an elongated structure, where the bases are positioned
extrahelically and the backbones occupy the center of the helix. This transition has only
been identified, however, at a degree of supercoiling substantially higher than is
generated physiologically.
To examine the structural changes resulting from physiological levels of positive
DNA supercoiling, I have developed a method for preparing highly purified positively
supercoiled plasmid substrates. Based on a method previously developed in this
laboratory, this allows for preparation of large quantities of very pure, highly positively
supercoiled plasmid. It also expands on earlier methods by exploiting ionic strength to
modulate the direction of supercoiling introduced, allowing preparation of either
positively or negatively supercoiled substrates.
A combination of approaches has been used to elucidate changes to DNA
structure that result from physiological levels of positive supercoiling. Enzymatic probes
for regions of single-stranded character are not reactive with positively supercoiled
plasmid, indicating that stably unpaired regions are not present. Additionally, the effect
of supercoiling on the activity of restriction enzymes has been examined. With the
enzymes tested, no substantial differences in cleavage rates were observed with either
positively or negatively supercoiled substrates. To examine structural changes at a wider
range of superhelical densities, design and preparation was undertaken on 2-aminopurinecontaining DNA substrates for use in fluorescence studies with a magnetic
micromanipulator. Technical limitations rendered these experiments infeasible with
current instrumentation, but important insights were gained for future fluorescence-based
micromanipulation experiments.
A destabilizing effect on the base pairs, however, can be seen using Raman
difference spectroscopy, suggesting a subtle shift toward the more extreme extrahelical
state. The Raman data suggest that structural adjustments due to positive supercoiling are
small but significant, and in addition to the base-pairing effects, alterations are observed
in phosphodiester torsion and the minor groove environment, as well as a slight shift in
sugar pucker conformation to accommodate lengthening of the DNA backbone.
These results point to subtle changes in DNA structure caused by biologically
relevant levels of positive superhelical tension and positive supercoiling. All of the
changes are consistent with the mechanical effects of helical overwinding and suggest a
model in which base pair destabilization in overwound DNA could affect the search
mechanisms used by DNA repair enzymes and the binding of other proteins to DNA.
Thesis Supervisor: Peter C. Dedon
Title: Professor of Toxicology and Biological Engineering
Acknowledgements
I would like to start by thanking my family, particularly my parents, Merritt and Jenny,
my grandmother Mary, my brother Stephen and my aunt Marita Jo Broadus for their
support. They never failed to offer encouragement when I needed it, and I couldn't have
done this without them.
The research experience I gained as an undergraduate was vital to my success in graduate
school, and for that I wish to thank my undergraduate thesis advisor, Professor George S.
Bailey, as well as Kate Mathews and Dr. Ulrich Harttig from his laboratory. They were
all amazingly kind and encouraging, and I appreciate their willingness to work with and
trust an undergraduate in a research environment.
I have gained a lot from my interactions with all current and former members of the
Dedon Lab, and I would like to extend my thanks to all of them. I would particularly like
to thank Debra Dederich, who has been invaluable in preparation of substrates for this
research, and whose dedication and hard work in the face of many technical challenges is
tremendously appreciated. I would also like to thank Dr. C. Eric Elmquist, who
performed the MS analyses of chloroacetaldehyde-treated plasmid samples, and Dr.
Michael DeMott who assisted greatly with the editing of this thesis. Dr. Koli
Taghizadeh, Yelena Margolin, Dr. Min Dong, and Dr. Christiane Collins all gave
important input into my research, and have been a pleasure to work with as well.
The support staff for the Dedon Lab has been absolutely amazing, and I would like to
thank them all. Marcia Weir, Kristine Marzilli and Dawn Erickson have all done much to
make my life easier, and have my gratitude. Olga Parkin is in a class all her own, with an
uncanny knack for solving any problem that might come up. Her assistance in all matters
administrative, as well as her friendship, have meant a great deal to me.
I would also like to thank several of the grad students and post-docs whose friendship I
have enjoyed through the years: Dr. Maxine Jonas, David Appleyard, Vasileios
Dendroulakis, Dr. Can Ozbal, Dr. Teresa Wright, Dr. Joe Newman, Dr. Maryann Timins,
Dr. Elaine Chin, Dr. Janice Lansita, Dr. Jane Sohn, and J.P. Cosgrove.
This thesis project has involved multiple collaborations, which have significantly
enhanced both my education and the quality of the information obtained. I would like to
thank Professor George Thomas at the University of Missouri at Kansas City for opening
up his lab to me for the Raman spectroscopic studies of supercoiling. Within the Thomas
lab, I wish to thank Professor James Benevides for sharing his expertise with me, and
allowing me so much instrument time on my visits to Kansas City. Additionally, he and
his family were wonderful hosts, and truly made me feel welcome during my stays there.
For their collaboration on the 2-aminopurine studies, I wish to thank Professor Peter So
and Dr. Serge Pelet for their input and experimental assistance.
I would like to thank my thesis committee, Professor Bevin Engelward, Professor John
Essigmann and Dr. Richard Roberts for all of their support and input. They've been
incredibly encouraging, and have been a wonderful source of challenging questions and
intriguing ideas.
For help in more ways than I can list, I would like to thank Gavin McNett. His keen
editorial eye and assistance with graphics have dramatically improved the quality of this
thesis. Beyond that, his kindness, good nature, wit, phenomenal cooking skill, and
(above all) patience have been vital to seeing me through the thesis writing process. I
find it difficult to adequately express my gratitude and respect.
Finally, I would like to thank my thesis advisor, Professor Peter Dedon, for all his support
and encouragement through the years. He has been incredibly generous and supportive,
and I truly appreciate him sticking with me through what proved to be a very challenging
and difficult project. His openness to new ideas and the freedom he has given me to
explore different approaches to the problems I have faced have greatly enriched my
education. I am truly grateful.
Biographical Note
Marita Christine Barth was born in 1975 in Dallas, Oregon, and graduated from Dallas
High School in 1993. After a summer internship at the Laboratory Services Division of
the Oregon Department of Agriculture, she enrolled at Oregon State University in
Corvallis, OR. Following a junior year abroad at Lincoln University in Canterbury, New
Zealand, she returned to complete a thesis project, entitled "In vitro Mechanisms of
Chlorophyllin Anticarcinogenesis Against Dibenzo[a, l]pyrene," in the laboratory of
Professor George S. Bailey. She was awarded a Bachelor of Science in Bioresource
Research with an option in Toxicology and a minor in Chemistry, in 1998. She
subsequently enrolled at MIT, and during the course of her graduate studies has been
awarded a National Defense Science and Engineering Graduate Fellowship, and a Poitras
Pre-Doctoral Fellowship.
Table of Contents
Abstract ....................................................... ........................
5
Acknowledgements ...................................................................
7
Biographical Note ....................................................................
9
List of Abbreviations ................................................................
15
List of Figures .........................................................................
17
Chapter 1 - Introduction
Packaging of DNA in Eukaryotic Cells ..................................
19
Topological Domains and DNA Supercoiling ............................
20
The Twin-Domain Model of DNA Supercoiling .........................
21
Mathematical Descriptions of Supercoiling .............................
23
Energetics of Supercoiling .................................................
26
...............
27
Physiological Roles for Negative Supercoiling ............................
30
Physiological Effects of Positive Supercoiling ..........................
34
Structural changes in negatively supercoiled DNA ......................
35
Structural Changes in Positively Supercoiled DNA .....................
37
Figures ........................................................................
41
Literature Citations .........................................................
47
Topoisomerases .........................................
Chapter 2 - Preparation of Positively and Negatively Supercoiled
Substrates
Introduction ...................................................................
55
Materials and Methods .........................................................
56
Results and Discussion ........................................................
60
11
Conclusions ....................................................................
63
Figures .................................................... ......................
65
Literature Citations .......................................
68
.............
Chapter 3 - Reactivity of Supercoiled DNA with Chemical and
Enzymatic Probes of DNA Structure
Introduction ....................................................................
71
Materials and Methods .........................................................
73
Results ........................................................................
76
D iscussion ......................................................................
81
Figures ............................................................
84
Literature Citations .......................................
89
.............
Chapter 4 - Raman Spectroscopic Studies of Positive Supercoiling
Introduction .....................................................................
93
Materials and Methods .........................................................
95
Results ........................................................................
96
D iscussion ......................................................................
100
C onclusions ....................................................................
105
Figures ........................................
106
Literature Citations ...........................................................
112
Chapter 5 - Design and Optimization of Fluorescent DNA Substrates for
Micromanipulator Studies
Introduction .....................................................................
115
Materials and Methods .........................................................
118
Results ........................................
120
D iscussion ................................................................
Figures ........................................
......
125
130
Literature Citations ...........................................................
135
Chapter 6 - Effects of Supercoiling on Cleavage by Type II Restriction
Enzymes
Introduction .....................................................................
139
Materials and Methods ......................................................
142
Results .........................................................................
143
D iscussion ......................................................................
146
Figures ........................................
149
Literature Citations ......................................
153
Chapter 7 - Conclusions
Conclusions .....................................................................
157
Literature Citations ...........................................................
164
Abbreviations
2AP
6MI
A
BCIP
bp
C
CBE
dA
dAPTP
dC
ALk
dG
DIG
DMS
dNTP
dT
DTT
EDTA
EA
EC
G
HPLC
IPTG
Lk
Lko
NBT
NMR
PCR
SDS-PAGE
Ua
T
TBE
TIR
Tris
Tw
Wr
2-Aminopurine
6-Methylisoxanthopterin
Adenine
5-Bromo-4-chloro-3-indolyl phosphate
Base pair(s)
Cytosine
Chicken blood extract
2'-Deoxyadenosine
2-Amino-2'-deoxyadenosine-5'-triphosphate
2'-Deoxycytidine
Linking number difference
2'-Deoxyguanosine
Digoxygenin
Dimethyl sulfate
2'-Deoxynucleotide triphosphate
2'-Deoxythymidine
Dithiothreitol
Ethylenediaminetetraacetic acid
1,N6 -Ethenoadenine
3,N4 -Ethenocytosine
Guanine
High pressure liquid chromatography
Isopropyl P-D-1 -thiogalactopyranoside
Linking number
Linking number of a completely relaxed molecule
p-Nitrotetrazolium blue
Nuclear magnetic resonance
Polymerase chain reaction
Sodium dodecyl sulfate-polyacrylamide gel electrophoresis
Superhelical density
Thymine
Tris-borate-EDTA
Total internal reflection
2-Amino-2-(hydroxymethyl)- 1,3-propanediol
Twist
Writhe
List of Figures
Figure 1.1:
Packaging of eukaryotic DNA into chromatin
Figure 1.2:
The twin-domain model of DNA supercoiling
Figure 1.3:
Twist and writhe parameters of DNA supercoiling
Figure 1.4:
Toroidal and plectonemic DNA writhe
Figure 1.5:
Formation of a "chicken foot" structure at a stalled replication fork
promoted by accumulated positive supercoiling
Figure 1.6:
Two general types of cruciform extrusion at inverted repeat sequences of
DNA
Figure 2.1:
Reaction scheme for preparation of supercoiled plasmid substrates
Figure 2.2:
Optimization and analysis of prepared supercoiled substrates
Figure 2.3:
Purification of closed-circular plasmid preparation by extraction with
acidic phenol
Figure 3.1:
Kinetics of reaction of S1 nuclease with supercoiled and relaxed substrates
Figure 3.2:
Reaction of S I nuclease with supercoiled and relaxed substrates
Figure 3.3:
Kinetics of reaction of nuclease BAL-31 with supercoiled and relaxed
substrates
Figure 3.4:
Reaction of chloroacetaldehyde with DNA bases
Figure 3.5:
Reaction of chloroacetaldehyde with supercoiled and relaxed substrates
Figure 4.1:
Raman spectra of negatively supercoiled and relaxed pUC 18 plasmid
DNA at 752 nm excitation
Figure 4.2:
Raman spectra of negatively supercoiled and relaxed pUC 18 plasmid
DNA at 532 nm excitation
Figure 4.3:
Raman spectra of positively supercoiled and relaxed pUC 18 plasmid DNA
at 752 nm excitation
Figure 4.4:
Raman spectra of positively supercoiled and relaxed pUC 18 plasmid DNA
at 532 nm excitation
Figure 4.5:
Raman spectra of negatively supercoiled and relaxed pUC 18 plasmid
DNA in D2 0
Figure 4.6:
Raman spectra of positively supercoiled and relaxed pUC 18 plasmid DNA
in D2 0
Figure 5.1:
Hydrogen bonding schemes of the canonical Watson-Crick adeninethymine base pair, the fluorescent base analog 2-aminopurine paired to
thymine, and a proposed protonated form of 2AP that could pair with
cytosine
Figure 5.2:
Preparation of components of micromanipulator substrate
Figure 5.3:
Assembly scheme for micromanipulator substrate
Figure 5.4:
Fluorescence lifetime of free and incorporated 2-aminopurine
Figure 5.5:
Comparison of original design of micromanipulator substrate with
optimized design
Figure 6.1:
Possible reaction modes of closed circular DNA with restriction enzymes
Figure 6.2:
Kinetics of reaction of NarI with supercoiled and relaxed substrates
Figure 6.3:
Kinetics of reaction of Ehel with supercoiled and relaxed substrates
Figure 6.4:
Kinetics of reaction of EcoRI with supercoiled and relaxed substrates
Chapter 1 - Introduction
Packaging of DNA in Eukaryotic Cells
The extensive length of genomes makes it necessary for both prokaryotic and
eukaryotic cells to compact their DNA. The human genome, for example, consists of
-3xl09 bp, and codes for an estimated 20,000-25,000 genes (1). For a single human
diploid cell, this amount of DNA laid end-to-end would result in a length of more than
two meters. Since the nucleus that must contain this DNA is a mere -10 -5 m in diameter,
a significant degree of compaction is necessary.
While complete compaction of the genome might be ideal for storage, as well as
for cellular events like chromosomal segregation, other cellular processes periodically
require at least some of the DNA to be unpackaged. RNA polymerases need to process
along the DNA to function, so any compaction would need to be removed from
transcriptionally active genes before transcription ensues. This means that in certain
stages of the cell cycle the DNA needs to be partially compacted and partially unpacked.
To further complicate the situation, cells need the ability to rapidly up- and downregulate genes in response to stimuli, so compaction must be both dynamic and tightly
regulated. All of this must be accomplished in a fashion that minimizes tangling and
knotting of the DNA molecules.
The eukaryotic solution to the problem is to store the DNA as chromatin, which
utilizes protein-DNA and protein-protein interactions to package the DNA in increasing
levels of compaction (reviewed in Ref. 2). First, the naked DNA is wrapped into
nucleosomes, in which a 147 bp segment of DNA is wrapped 1.8 times around a core of
positively charged histone proteins in a left-handed toroid. The nucleosomes are
separated from one another by 50-100 bp segments of linker DNA, giving extended
nucleosomal arrays the appearance of "beads on a string" when viewed in electron
micrographs. This nucleosomal array, also known as the 10 nm fiber, is further
compacted into a secondary structure known as the 30 nm fiber, although debate persists
as to whether this takes a solenoidal (3,4), or irregularly zig-zagging "slinky-like" form
(5,6). Further compaction of chromatin into a tertiary structure requires long-range
interactions of the chromatin fiber, and it is affected by a number of different proteins in
a process that is still not well understood (Fig. 1.1).
Topological Domains and DNA Supercoiling
One important consequence of the compaction into chromatin is the division of
DNA molecules into topological domains (7). The DNA in chromatin is anchored to the
nuclear matrix at intervals, in such a fashion that it cannot freely rotate. The DNA
between any two of these points is known as a topological domain. In an unconstrained
linear DNA molecule, any change in twist that is introduced will diffuse through the
molecule and will gradually dissipate, as one strand of the DNA rotates relative to the
other. When such free rotation is limited, as happens within a topological domain, any
added twist will be retained by the DNA, since it cannot diffuse past the boundaries of
the domain. Topological domains vary dramatically in size. In human cells, the most
recent estimate ranges from <95,000 to >150,000 bp (8), although other estimates offer a
broader distribution from 1,000 to 300,000 bp for eukaryotic genomes (9-11). If an
average of 100,000 bp/domain is assumed, this would suggest that a diploid human cell
has -60,000 topological domains.
Supercoiling in DNA occurs when the DNA is either underwound (negative
supercoiling) or overwound (positive supercoiling) relative to the B-form helix.
Constrained toroidal supercoiling, in the form of DNA wrapped around histone cores, is
present in cells independent of any topological domains. This supercoiling is trapped by
the binding of the proteins to the DNA, and as such cannot diffuse out. Because DNA
wraps around histones in a left-handed fashion, the supercoiling that is constrained is
negative.
The partitioning of cellular DNA into topological domains does allow the
retention of unconstrained supercoiling. This type of supercoiling is free to diffuse
through the molecule, within the limits set by the boundaries of the topological domain.
This normally occurs when the strands of DNA are separated during various cellular
processes. The earliest detailed description of this phenomenon is the twin-domain
model of transcription proposed by Liu and Wang (12).
The Twin-Domain Model of DNA Supercoiling
In the twin-domain model, Liu and Wang account for the consequences of the
requirement for the RNA polymerase and the DNA template to rotate relative to each
other during the transcription process (12). In theory, either molecule could rotate,
however evidence indicates that polymerases are anchored to the nuclear matrix (13,14).
Therefore, for transcription to occur, the DNA must rotate relative to the polymerase. As
this happens, negative supercoiling is generated behind the complex, while positive
supercoiling is generated ahead of it. (Fig. 1.2) This supercoiling can diffuse rotationally
along the helix, but cannot diffuse past the limits of the topological domain.
According to this model, the degree of supercoiling present in a topological
domain is determined by the orientation of transcribed genes, by how transcriptionally
active these genes are, and by the activity of different topoisomerases within the cell. If
two transcribed genes are oriented in the same direction, the supercoils will eventually
cancel each other in the region between the genes, as the positive supercoils generated
ahead of the one gene are met by the negative supercoils generated behind the other. If
the genes are oriented in opposite directions, significant supercoiling of either sign can be
generated between the two genes, as it will be accumulating from both directions.
Topoisomerases can affect the degree of supercoiling within a topological domain by
selectively adding or removing supercoils. If a topoisomerase that is removing
supercoiling of one sign is more active than the topoisomerase removing the other, an
imbalance will develop in the net amount of supercoiling. Topoisomerases will be
discussed in greater detail later in this chapter.
Evidence in support of the twin-domain model has been found in vitro (15,16), in
prokaryotes (17,18), and in eukaryotes (19,20), illustrating its importance as a widespread
biological phenomenon. The phenomenon also isn't limited to transcription. While the
original model described transcriptionally-induced supercoiling, any protein that forces
rotation of the DNA template can generate supercoiling in a similar fashion. This
includes helicases such as the E. coli nucleotide excision repair complex UvrAB (21) and
the SV40 large tumor antigen (22), and translocating enzymes such as the type I
restriction enzyme EcoAI (23).
Mathematical Descriptions of Supercoiling
Before delving into the structural and physiological consequences of supercoiling,
it is useful to consider the mathematical conventions used to describe it. In most
respects, a topological domain is analogous to a closed-circular DNA molecule, so
several of the same concepts apply.
DNA has a double helical structure comprised of two interwound strands. The
linking number of the DNA, Lk, is the number of times the strands are intertwined for a
given molecule. For B-form DNA, the linking number of a completely relaxed molecule,
Lko, can be obtained by dividing the number of base pairs, n, by the helical repeat. In
physiological solution conditions, the helical repeat is -10.4 bp/turn (24), so:
Lko = n/10.5
After torsion has been introduced into the molecule, Lk will change by 1 for every
complete turn of the helix added (or removed). The linking number difference, ALk, is
equal to the new linking number minus the relaxed linking number:
ALk= Lk- Lko
In a closed-circular molecule, both strands of the DNA are unbroken, which means that
Lk must be an integer value. From the definition of Lko, however, we can see that it is
not necessarily an integer value. Thus, depending on the precise length of the DNA,
some closed-circular molecules of DNA cannot reach a completely relaxed state. It also
means that ALk is not necessarily an integer value. Because changes in ALk for an intact
molecule can only occur by breaking one or more strands of the DNA and then
introducing full turns before religation, changes in ALk must be of an integer value.
It is also important to note that under consistent solution conditions, localized
supercoiling can occur in an intact topological domain, but the net Lk for that domain
cannot change without breakage of at least one strand of the DNA followed by religation. This means that if one portion of an intact molecule is subjected to a localized
positive linking number change, a compensatory negative supercoiling of equivalent Lk
value must occur elsewhere in the domain.
Two closed circular molecules differing only in ALk are referred to as
topoisomers. Populations of topoisomers can be resolved using agarose gel
electrophoresis, as an increase in IALkJ will result in increased mobility of a plasmid on a
gel (25).
Changes in the linking number describe differences between topoisomers of the
same molecule, but give no information about the length of DNA that is accommodating
the introduced torsion. A ALk of +1 would put considerably more strain on a 100 bp
molecule than it would on a 1,000 bp molecule. To account for the length of a molecule
when describing supercoiling, the superhelical density, o, is used:
a = ALLk/Lko
Superhelical density values allow for comparison of the degree of supercoiling among
molecules of different sizes. DNA isolated from most cells is negatively supercoiled; in
E. coli the superhelical density will generally vary from o Z -0.03 to -0.09, depending on
growth conditions (26-28).
Since the linking number represents the number of times the two strands of the
DNA are intertwined, it is tempting to think of changes in Lk as simple changes in the
twist (or helical repeat) of the molecule. However, this is oversimplified. By way of
analogy, consider an electrical cord that has one end held stationary while the other end is
turned. As the number of turns increases and twists along the length accumulate, the cord
will eventually begin to fold upon itself. DNA behaves in a similar fashion; Lk is
partitioned between two components - twist and writhe:
Lk= Tw + Wr
The twist value represents the number of times the strands wrap around the helical axis,
while writhe is the crossing of the helical axis over itself (Fig. 1.3). The double helical
structure of DNA means that twist is always present, while writhe will only occur when
Lk • Lko. Therefore:
ALk = ATw + Wr
Because of this relationship between twist and writhe, any change in one will
necessarily affect the other. Partitioning between the two can therefore be changed by
any factor that affects either component. A number of factors have been identified that
can shift this distribution.
A change in twist effectively means a change in the helical repeat of the DNA.
With underwinding, the helical repeat will be >10.4 bp/turn, while with overwinding it
will be <10.4 bp/turn. Intercalating agents are one example of a factor that will change
the twist component of DNA. They act by introducing localized unwinding of the DNA
at the site of intercalation, which requires a compensatory change in the writhe
component of the DNA. Depending on the degree of intercalator binding, the writhe may
actually have to be of the opposite sign, with a positive writhe offsetting an excess of
negative twist.
Writhe exists in two forms, toroidal and plectonemic. Toroids form when the
DNA is wrapped around a central axis such as a histone complex. Unconstrained
supercoiling, such as occurs in the absence of toroidal wrapping in nucleosomes, takes a
plectonemic form (29,30) (Fig. 1.4). Salt conditions have a significant effect on
plectonemic writhing in DNA. The highly negatively charged backbone of DNA makes
it difficult for different segments of the molecule to come into close proximity during the
crossing-over that occurs in plectonemic writhing. Cations in the salt solution help to
neutralize this charge repulsion, reducing the effective diameter of the DNA and allowing
it to interwind more tightly (31). At physiological salt conditions, writhe has been
estimated to account for approximately two-thirds to three-quarters of the linking number
difference (29,30,32).
Energetics of Supercoiling
A DNA molecule is considered to be supercoiled if it has a ALk # 0 and there is a
considerable free energy cost associated with supercoiling. The free energy of
supercoiling (AG,,) has been empirically determined by measuring the binding affinities
of intercalating dyes (33), and examining distribution of topoisomers (34,35). It has also
been estimated computationally, using Monte Carlo simulations (32). The results are in
general agreement (for a comparison, see ref. (32); the dependence of free energy on the
degree of supercoiling is quadratic:
AGsc=KRT(ALk) 2/N
Where R is the gas constant, T is the absolute temperature, N is the number of base pairs,
and K is a length- and ionic strength-dependent constant. As ionic strength increases, K
decreases, resulting in a reduction in the free energy of supercoiling (35). The length
dependence of K is more significant for shorter segments of DNA; above -2000 bp K is
inversely proportional to N, and the value of NK becomes length independent (36).
The enthalpy (AH) and entropy (AS) of supercoiling have been estimated
computationally by Monte Carle simulations (37), and also measured experimentally
using microcalorimetry (38), Gibbs-Helmholtz (39), and van't Hoff (40) methods.
Experimental methods show the enthalpy of supercoiling to be positive, and substantially
larger than the free energy. Given this, and the relationship:
AG = AH - (TAS)
We can conclude that the entropy of supercoiling must also be large and positive. This is
in direct contradiction to the computationally predicted values, which calculate a negative
entropy for supercoiling. Intuitively, a negative entropy makes sense: The limitations in
the conformations which the DNA can adopt upon supercoiling would result in a negative
entropy for supercoiling. This is not experimentally borne out, however, so the small
decrease in configurational entropy must be more than compensated for by a much larger
positive entropy component. This has been hypothesized to be due to the disruption of
bound ions and water molecules resulting from the twisting and bending motions of the
DNA upon supercoiling (32), as well as changes in local interactions between base pairs
and between base pairs and solvent (39).
Topoisomerases
Topoisomerases are the enzymes responsible for regulating and maintaining
levels of supercoiling within a cell. They can be classified into two major types, I and II.
Type I enzymes function via a swivel mechanism, cleaving one strand of the DNA and
either allowing or driving rotation prior to re-sealing the cleaved strand (41,42). Type II
topoisomerases act via a strand passage mechanism: Both strands of the DNA are cleaved
and held open by the enzyme, like a gate, while another segment of the molecule is
passed through the opening (43). The mechanisms are directly related to the degree to
which the topoisomerases change the linking number of the molecule. Type I enzymes
can alter ALk in single integer increments, while Type II always change the linking
number in multiples of two.
Both major types of topoisomerase are further split into two sub-families, A and
B. Type lA enzymes pass the intact strand through the gap created by the single strand
cleavage prior to religation, changing the linking number in steps of __l (44). While most
lA enzymes require only a metal as a cofactor, reverse gyrase is a unique type lA that
can introduce positive supercoils in an ATP-dependent fashion (45,46). Type 1B
enzymes, while also cleaving a single strand, are thought to allow rotation around the
intact strand, changing the linking number by an average of as much as ±5 in a single step
(47).
In most known cases, Type I topoisomerases act to remove superhelical tension
from the molecule. In large molecules, however, thermal fluctuations in structure can
occur that result in a molecule being apparently relaxed at the site of topoisomerase
action while there is still some supercoiling present in the molecule. Because of this,
whenever a Type I topoisomerase acts to relax a population of supercoiled molecules, a
distribution of topoisomers, rather than an entirely relaxed population, will result. The
pattern of topoisomer distribution is predictable and follows a Boltzmann distribution,
with the median at the most relaxed possible state for the given conditions (34).
Type II topoisomerases utilize ATP to drive their reaction, and depending on the
specific enzyme, can either relax DNA or introduce supercoils. Types IIA and IIB both
exhibit similar mechanisms, with the primary difference between the two sub-families
being global structural features. These enzymes change the superhelicity of a molecule
using a sign inversion mechanism: At the site of a DNA crossover, when both strands of
a molecule are cleaved and another segment of the molecule is passed through, it reverses
the handedness of the crossover (e.g changing a positive writhe to a negative writhe),
resulting in a change in ALk of ±2 (48). Because of the double-stranded nature of the
break induced by Type II topoisomerases, under certain conditions they can
catenate/decatenate and knot/unknot DNA in addition to their function in modulating
superhelicity (49,50).
While the mechanism and degree of supercoiling change is defined by the
subfamily to which a topoisomerase belongs, the directionality of the supercoiling that is
removed (or introduced) varies with the enzyme. Some topoisomerases will only act on
supercoils of a specific handedness, while others will remove supercoils of either sign.
The balance of activities of these different topoisomerase species controls the overall
level of unconstrained supercoiling within a cell.
One well-studied example of this topoisomerase balance is in E. coli, where the
level of supercoiling is tightly homeostatically regulated (51,52). It is primarily
controlled by two enzymes: The type IA enzyme topoisomerase I is responsible for
removal of negative supercoils (53), and the type II enzyme gyrase selectively removes
positive supercoils and introduces negative ones (54,55). Increasing the level of negative
supercoiling results in an up-regulation of topoisomerase I expression and a downregulation of gyrase (56); reducing the level of supercoiling has the opposite effect (51).
Perturbation of the activity of either of these enzymes through gene alteration (53) or use
of chemical inhibitors (57) will change the overall level of supercoiling in the cells. Until
recently, a deletion of the topA gene coding for topoisomerase I was thought to be lethal
without compensatory gyrase mutations (58). Strains have now been created where this
is not the case, although deletion of topoisomerase III in addition to topA will result in
non-viable cells (59).
Topological controls are also present in eukaryotic cells (19,60,61), suggesting
that this phenomenon is of broad biological relevance. Taken together, this evidence
suggests that tight control of supercoiling is important for cellular survival, and as such,
superhelicity must play a vital physiological role.
Physiological Roles for Negative Supercoiling
The importance of supercoiling implied by these tight controls bears out, with
many cellular processes are affected by supercoiling. Physiological effects can arise
from the alteration in either the twist or the writhe of the molecule. Effects due to twist
can result from the change in the helical repeat, transitioning to alternative secondary
structures, and torsional tension on the helix. Writhe is required for compaction,
facilitates looping and bending, and can bring distant elements on a single molecule into
close proximity. One area that has been well studied in regards to supercoiling is gene
expression; the supercoiling state of the molecule can affect gene expression via a
number of different mechanisms.
One way in which negative supercoiling can affect gene expression is by assisting
in open complex formation in the DNA (62). When the RNA polymerase complex
initiates transcription, it first binds to the -35 and -10 elements in the promoter, forming a
closed complex (63). This is followed by nucleation, where DNA in the -10 region is
unwound. The open region of the DNA is then extended to the start site of transcription,
allowing the polymerase complex to initiate transcription (reviewed in (64). Denaturation
of DNA is energetically unfavorable, and creation of an open region of DNA by a protein
carries a significant energetic cost. When the DNA is negatively supercoiled, this
energetic cost can be "paid" by the supercoiling-associated free energy trapped in the
molecule, concentrating the negative supercoiling into the denatured region and as a
result relieving the superhelical strain on the remainder of the molecule (65).
While a high degree of negative supercoiling is useful for stabilizing denatured
regions of DNA, other mechanisms that control gene expression are more precise in the
degree of superhelical density required to exert an effect. Increasing the degree of
negative supercoiling will not always increase the level of expression of a particular gene.
Early evidence of this was found by Stirdivant and co-workers, who examined two
adjacent genes from the maize chloroplast chromosome and discovered that they were
optimally expressed at different levels of supercoiling in vitro (66). Interestingly, while
one gene exhibited an increase in expression followed by a plateau as negative
supercoiling increased, the other showed an increase, peaked, and then decreased under
the same conditions. This suggests that multiple mechanisms are responsible for the
supercoiling-based changes in expression levels.
Steck and co-workers did a broader study in vivo and discovered that the optimal
level of supercoiling for gene expression in E. coli could generally be correlated to the
length of the spacer DNA between the -10 and -35 hexameric elements in the promoter
region of the genes (67). Genes with a 17 bp spacer were optimally expressed at normal
levels of supercoiling. For genes with a shorter spacer, a reduction in negative
supercoiling increased expression, and for those with a long spacer, increased levels of
negative supercoiling increased expression.
The spacer phenomenon can be explained in terms of the relative rotational
orientation of the -10 and -35 elements on the molecule (68). The RNA polymerase
complex binds to both elements during initiation, and as such the rotational orientation of
the elements is an important determinant of binding. If 17 bp is an optimal spacer for the
elements to be in a given orientation at physiological levels of supercoiling, then a longer
spacer would need an increased level of negative supercoiling to have the same number
of helical turns (and hence the same orientation) in between the two elements. The
reverse is true for a shorter spacer.
The writhe component of DNA supercoiling can also play a role in gene
expression. One notable example of how it may do so involves the ability of prokaryotic
enhancers to affect transcription when they are located at a substantial distance from the
promoter. This works by increasing the probability that the enhancer will be brought
close enough to the promoter for the two to interact. In a molecule with no writhe, any
effort to bring the two elements together is essentially a search in three-dimensional
space. Normal fluctuations in the molecule may bring the two elements into proximity,
but since the chances of the two elements meeting in three-dimensional space are low,
these events are infrequent. When the molecule is crossed over itself, as in writhe, the
search becomes two-dimensional; the molecule can move along itself in a slithering
fashion (reptation), and in this manner dramatically increase the odds of bringing two
distant sites into close proximity. A Monte Carlo simulation estimated that for sites
separated by 3 kb, the probability of the two sites coming into contact is two orders of
magnitude higher when the DNA is subjected to physiological levels of supercoiling than
when it is relaxed (37). Experiments in vitro with the glnAP2 promoter and the NtrCdependent enhancer from E. coli confirm this prediction: in a molecule where the
promoter and enhancer are separated by 2500 bp, effective communication between the
two elements is increased 50-fold when the molecule is negatively supercoiled relative to
when it is relaxed (69). This observed difference was significantly reduced when the
promoter and enhancer were located closer together on the molecule.
Control of gene expression by negative supercoiling is not limited to these
mechanisms, nor are the physiological effects of negative supercoiling limited to control
of gene expression. In addition to affecting the initiation of transcription, negative
supercoiling has been demonstrated to play a role in transcription elongation, as well
(70). In other cellular processes, the twist and the writhe components of supercoiling
have been implicated in the binding of histone proteins to DNA (71), replication (72),
site-specific recombination (73), and transposition (74).
Physiological Effects of Positive Supercoiling
The discussion thus far has been limited to the effects of negative supercoiling.
However, evidence suggests that positive supercoiling has important physiological
consequences as well. These have not been as well elucidated; plasmid DNA isolated
from eubacteria under normal conditions is negatively supercoiled, making it a much
more easily obtained substrate than positively supercoiled plasmid, which must be
prepared artificially. Additionally, because DNA isolated from most organisms is
negatively supercoiled, it is a more obvious candidate for study. Still, the twin-domain
model predicts the existence of at least transient positive supercoiling in cellular DNA,
and this presence has been verified experimentally (17,19,75). Also, several proteins
have been identified that can introduce either free or constrained positive supercoiling
into DNA independent of a helix-tracking process, including the archaebacterial
topoisomerase reverse gyrase (46), and the ubiquitous human chromatin-associated
protein DEK (76). This suggests an important role for positive supercoiling in the
functioning of the cell.
With the enhancing effect of negative supercoiling on transcription and
replication, an obvious function of positive supercoiling would be to act as a negative
regulator of these processes. Any process that requires unwinding of the helix will face a
growing energy barrier to activation as the degree of positive supercoiling increases.
Positive supercoiling has been demonstrated to inhibit transcription in vitro at levels as
low as a = +0.03, potentially acting by interfering with initiation and elongation steps
(77). Histone complexes bind with less affinity to positively supercoiled DNA (71,78),
leading to the theory that waves of positive supercoiling generated ahead of helix-
tracking enzymes can help to drive nucleosomes off the DNA, making it more accessible
(71,78). Accumulated positive supercoiling can force regression of a stalled replication
fork to form a "chickenfoot" structure analogous to a Holliday junction (Fig. 1.5) (79).
Most recently, evidence has been found that positive supercoiling significantly enhances
the rate of telomere resolution in the linear chromosomes of the Lyme disease spirochete
Borrelia burgdorferi(80). The diversity of processes that appear to be affected by
positive supercoiling suggests a broad biological role, but considerably more information
is necessary to get a more complete picture of its effects within a cell.
Structural changes in negatively supercoiled DNA
Since plasmid DNA isolated from eubacteria under standard growth conditions is
negatively supercoiled, a ready substrate is available for researchers to use in examining
the effects of helical unwinding on the structure of DNA, and the phenomenon is wellstudied. Negative supercoiling increases the susceptibility of DNA to single-strand
specific nucleases (81-83), a result that has been attributed to stabilization of regions of
denatured base pairs within the DNA. The nature of negative supercoiling lends itself to
denaturation; the two strands of the DNA molecule become less intertwined, making
disruption of base pairing more likely as the level of underwinding increases. This
phenomenon has been treated theoretically by Benham, who predicted that stably
denatured regions would occur at physiologically expected levels of untwisting
(equivalent to c•
-0.05) (65). As the level of negative supercoiling increases, so too
does the likelihood of forming a denatured region.
In some cases, certain sequences of DNA can undergo more substantial transitions
to alternative secondary structures as a result of negative supercoiling. These alternative
structures occur in much the same way as stably denatured regions, with the free energy
that is stored in the molecule as supercoiling acting to stabilize otherwise unstable
conformations. This concentration of free energy into a relatively small region of the
DNA molecule reduces the superhelical strain in the rest of the topological domain. One
example of this phenomenon is cruciform extrusion, which occurs at sites of inverted
repeats in the DNA.
Cruciforms are branched DNA structures in which an inverted repeat, the halves
of which are usually separated by a short segment of non-palindromic sequence, extrudes
from the helix to form two opposing stem-loop structures (84). Cruciform extrusion can
happen by one of two processes. In C-type extrusions, the formation of the cruciform is
preceded by the stable melting of the palindromic region of DNA, while S-type
extrusions are more complex and involve the formation of partially extruded
intermediates that develop into cruciforms through a branch migration process (Fig. 1.6).
The mechanism of extrusion is dictated by the sequence context of the inverted repeat;
the sequences flanking the ColE 1 inverted repeat, a prototype for C-type extrusion, are
A-T rich and more likely to be prone to denaturation. Consistent with the requirement for
large-scale melting, C-type cruciforms extrude preferentially under low salt, and have a
high activation energy. S-type cruciforms, which are much more common, require salt of
at least 50-60 mM concentration and have a lower activation energy for extrusion
(85,86).
There is still some debate over the physiological role of cruciforms. Some studies
have suggested that such structures can affect gene expression, forming above a certain
negative superhelical density and acting as a block to the progression of the polymerase
complex (77). The structure has also been demonstrated to be a specific binding site for
regulatory proteins. In one instance, a combination of cruciform extrusion and the
interaction of the cruciform with the RuvA protein from E. coli appears to act as a
topological switch, regulating processes that require interaction of cis-acting DNA
elements by controlling the reptation of supercoiled molecules (87).
Besides denatured regions and cruciforms, negative supercoiling can assist with
the formation of other alternative structures as well. These include left-handed Z-DNA
(88) and triplex H-DNA (89), both of which have their own unique sequence and
supercoiling requirements, and their own predicted biological function(s).
Structural Changes in Positively Supercoiled DNA
As is the case with physiological effects, much is unknown about the structural
consequences of positive supercoiling in DNA. Again, since positively supercoiled
substrates are much more difficult to obtain than the negatively supercoiled form, they
have not been studied in as much depth. Most studies are limited by a low level of
positive superhelical density in the substrates, or by a reliance on the presence of small
molecules bound to the DNA helix to generate positive supercoiling. Much of the current
information about structural consequences of positive supercoiling comes from
computational studies, and suffers from a lack of supporting experimental evidence.
Understanding the structural changes that are induced by positive supercoiling is vital to
further understanding the mechanisms of the physiological effects already hypothesized,
as well as predicting effects that have not yet been identified.
An important insight into structural changes caused by positive supercoiling came
from Allemand et al. (90). This group used single-molecule biomechanical experiments
to look at global structural changes in DNA at varying levels of introduced torsion. A
long (17,000 bp) piece of DNA was attached at one end to a solid support, and on the
other to a paramagnetic bead. Magnets were used to rotate the bead, introducing precise
amounts of either positive or negative torsion into the molecule. The magnets were also
used to stretch the DNA, preventing writhe, and therefore forcing the entire ALk to be
partitioned into twist. Both the length of the DNA and the force being used to pull on the
bead can be measured using this system. To monitor changes in DNA structure resulting
from introduced torsion, both length vs. force at a constant o, or length vs. oy at a constant
force were measured.
In the case of underwound DNA, the molecule underwent denaturation as a means
of relieving the overall torque (90). This phenomenon has been previously established
through modeling (65), experiments with negatively supercoiled plasmids (39,91), and
micromanipulator studies (92). With overwinding of the DNA, a different structural
transition occurs, albeit at a higher level of force (3 pN vs. 0.3 pN for underwinding).
Modeling of the data led to a proposed structure with a helical repeat of 2.6 bp/turn where
the DNA is significantly (-75%) longer than B-DNA, and the phosphate backbones
occupy the center of the helix, with extruded bases along the outside. Similarities to the
DNA structure originally proposed by Linus Pauling (93) led to the structure being
termed "P-DNA" (90). The solvent-exposed positioning of the bases was confirmed by
reaction of the twisted molecules with glyoxal, a chemical agent that reacts selectively
with unpaired bases (94).
In both under- and overwinding, the structural change occurs as a phase transition,
where above a certain critical force (and associated critical torque) writhing becomes
energetically unfavorable (90). At this point, the molecule adopts regions of the
alternative structure to alleviate strain. Just as in the case of cruciform structure
discussed previously, a small region of alternative structure can reduce the strain on the
rest of the molecule considerably; in the case of overwinding, adopting the new structure
reduces the twist in the rest of the molecule by three turns for every 10.4 bp converted.
While evidence for a non-base paired structure is compelling, much of the study
was completed under conditions that would not be expected physiologically.
Experiments were carried out at low salt (10 mM phosphate), at which the helix is less
stable than at physiological ionic strength; experiments in higher salt showed that greater
thresholds were necessary to observe the same structural transitions. Additionally, the
entire linking number difference was confined to twist, a partitioning that would not be
expected in unrestrained supercoiling. Conditions resulting in this phase transition may
be transiently generated immediately ahead of a processing helix tracking enzyme, but
are less likely to be sustained in other circumstances.
At the same time, it may be possible for some degree of base unpairing to be
present at lower levels of torsional strain. In negatively supercoiled plasmid, single
strand specific nuclease sensitivity can be detected at values as low as c = -0.02 (82),
despite the denaturation phase transition requiring significant pulling force to occur at a
c = -0.015. Could some regions of P-DNA-like structure develop in DNA that is
overtwisted to a lesser extent than the critical force that was identified in the
micromanipulator experiments?
There is some experimental evidence to suggest this may be possible. Nuclease
BAL-31 reacts with DNA when enough of the intercalating agent ethidium bromide has
bound to give an estimated superhelical density of Ya +0.15 (82). A long AT(,n) tract
becomes sensitive to the unpaired-base specific agent osmium tetraoxide when positive
supercoiling is induced by actinomycin D, a CpG binding agent (95). While these results
are intriguing, both studies suffer from a major flaw: reliance on a bound molecule to
introduce positive supercoiling prevents a realistic assessment of how positively
supercoiled DNA behaves in the absence of a bound ligand.
The ability to prepare positively supercoiled substrates with a high superhelical
density in the absence of extraneous ligands is crucial for further understanding of the
structural consequences of positive supercoiling in DNA. In experiments presented in the
succeeding chapters, I describe modifications to such a method that was previously
developed in this laboratory that improve the consistency and purity of prepared
positively supercoiled substrates. Using these substrates, I have performed chemical,
enzymatic, biomechanical, and spectroscopic studies to characterize structural changes
that occur with positive supercoiling. Taken together, these studies should provide an
important guide for future research into the structural and physiological consequences of
positive supercoiling in DNA.
~0~0~
DNA is wraps around histone
complexes to form
nucleosomes
Nucleosomes are compacted
into the 30 nm fiber
Ail
Chromatin loops attach to
protein matrix to form
topological domains
Further compaction leads to
fully condensed chromosome
Figure 1.1. Packaging of eukaryotic DNA into chromatin. DNA is packaged in successive
stages of compaction, going from naked DNA through nucleosomes and the 30 nm fiber to
chromatin loops, and finally a completely condensed chromosome.
Negative Supercoiling
(underwinding)
> 10.4 bp/turn
Positive Supercoiling
(overwinding)
< 10.4 bp/turn
X%^A#4%9
je helix-tracking enzyme
as RNA polymerase)
Figure 1.2. The twin-domain model of DNA supercoiling. As a helix-tracking enzyme
moves along a DNA molecule, the two must rotate relative to each other. If the enzyme is
prevented from rotating, the DNA molecule itself must rotate, causing a change in the helical
repeat of the DNA (supercoiling). Positive supercoiling (overwinding) will be generated
ahead of the enzyme, while negative supercoling (underwinding) will be generated behind it.
If the ends of the DNA are constrained in such a fashion that they cannot freely rotate, this
supercoiling will be retained in the DNA.
underwinding
overwinding
B
Figure 1.3. Twist and writhe parameters of DNA supercoiling. A) Twist refers to the number
of times the strands wrap around the helical axis. Supercoiled DNA can be either under- or
over-twisted relative to relaxed B-DNA B) Writhe is the crossing of the DNA helical axis
over itself. The direction of cross-over is determined by the direction of suDercoiline.
A
B
K~)
Figure 1.4. Toroidal and plectonemic DNA writhe. A) Toroidal writhe occurs when the DNA
wraps around a central axis, such as a protein. B) Plectonemic writhe is unconstrained and
occurs in the absence of toroidal wrapping.
A
Figure 1.5: Formation of a "chicken foot" structure at a stalled replication fork promoted by
accumulated positive supercoiling. A: A lesion on the leading strand causes replication to
stall. Arrowheads indicate the 3' ends of the strands; parent strands are depicted in black
while daughter strands are in red. B: A regression of the stalled replication fork promoted by
accumulated positive supercoils ahead of the fork. The daughter strands anneal to each other
due to this regression, forming a "toe" in the "chicken foot". Figure adapted from reference
74.
1111111 lllilllllIl1illnllilnlrir
A
!1111:!111
Figure 1.6: Two general types of cruciform extrusion at inverted repeat sequences in DNA.
A. S-type extrusion: Extrusion is initiated by formation of a small denatured bubble that
develops into opposing hairpin loops. Branch migration leads to a fully extruded cruciform
structure. B. C-type extrusion: under low salt conditions a large denatured bubble forms.
This denatured region form the full cruciform directly. Negative supercoiling is necessary for
either type of extrusion; the type a particular sequence undergoes is sequence dependent.
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Chapter 2 - Preparation of Positively and Negatively Supercoiled
Substrates
INTRODUCTION
The presence of superhelical tension in DNA is ubiquitous in the genomes of both
prokaryotes and eukaryotes (1-4), and is critical to of the physiology of cellular DNA (510). However, comparative studies of the physical and biological chemistry of different
supercoiling states of DNA are hampered by the lack of methods to produce the negative,
relaxed and positive states of DNA supercoiling in large quantity. While negatively
supercoiled plasmid is readily available for study (plasmid isolated from E. coli typically
has a superhelical density of -0.06), positively supercoiled DNA is not as readily
obtained. Furthermore, modified bases and base analogs cannot readily be incorporated
into supercoiled DNA, much less into DNA with different states of supercoiling. In order
to obtain substrates for my studies, I have developed a method for the preparation of
DNA in the three states of supercoiling, thus providing biochemically well-controlled
DNA substrates for comparison of different states of superhelical tension.
Several strategies exist for obtaining supercoiled molecules, including interfering
with the topoisomerase balance within growing cells (11), treatment of closed-circular
molecules with enzymes that introduce supercoils, such as gyrase or reverse gyrase
(12,13), and binding of the DNA with an agent that will constrain supercoils followed by
removal of compensatory free supercoils (14,15). While some (though by no means all)
of these methods fall short with regards to ease of preparation, cost of production, and/or
level of introduced superhelical density, they are all lacking in versatility; none allow for
introduction of either positive or negative supercoils using a single method.
Using the method for preparing positively supercoiled plasmid developed
previously by this lab (15) as a basis, I have developed a modified protocol to prepare
either negatively or positively supercoiled plasmid with a high superhelical density from
relaxed, closed circular DNA. This method exploits an observation that ionic strength
determines the direction of DNA binding to the archaeal histone rHMfB (16), using salt
conditions to control the sign of the supercoils induced in the plasmid substrates.
MATERIALS AND METHODS
Preparationofplasmid DNA. Plasmid pUC18 DNA was isolated from
transformed E. coli strain DH5a using a standard alkaline lysis procedure. Prior to use,
10 mg quantities of DNA in 5 ml of TE (10 mM Tris. 1 mM EDTA; pH 8.0) were
dialyzed in 100,000 MWCO dialysis tubing against 1 change of 4 L of 3 M NaCl in TE,
followed by 4 changes of 4 L of TE alone.
Preparationof topoisomerase-containingextract. A topoisomerase-containing
extract of chicken erythrocyte nuclei (herein referred to as chicken blood extract or CBE)
was prepared from sterile, citrate-treated chicken blood (Rockland Immunochemical,
Gilbertsville, PA) according to the method of Camerini-Otero and Felsenfeld (17). We
routinely process 200 mL of chicken blood in a single batch; the resulting extract is
aliquoted and stored at -800 C. One unit of CBE was defined as the amount of the extract
required to completely relax 1 pig of pUC18 under standard relaxation conditions (see
below). The nicking activity of the CBE (i.e., endonuclease activity as opposed to
topoisomerase activity) was assayed by resolving portions of the relaxed plasmid on 1%
agarose gels containing 0.25 [tg/mL ethidium bromide such that relaxed closed-circular
plasmid DNA was separated from single-strand nicked plasmid DNA. Batches of CBE
that showed significant (>25%) nicking at a ratio of 10 units of CBE per ýpg of plasmid
were discarded. Most batches of CBE caused nicking of less than 20% of the supercoiled
pUC18 plasmid (data not shown). Inactivated CBE for use in preparing sham-treated
substrates was prepared by heating CBE to 95 OC for 20 min, which eliminated all
nicking and toposiomerase activity.
Isolation ofrHMfB. An rHMfB-expressing strain of E. coli was created by
transforming electrocompetent JM105 with plasmid pKS323 (generously provided by K.
Sandman and J. Reeve, Ohio State University). rHMfB was isolated as described in (18),
with several minor changes. Cells from IPTG-induced cultures were lysed by two
passages through an Emulsiflex-C5 homogenizer (Avestin, Ottawa, ON) at 20,000 psi.
Affinity chromatographic purification of rHMfB was performed on a Hi-Prep 16/10
Heparin FF column (GE Healthcare, Piscataway, NJ) with buffer and gradient conditions
as described elsewhere (18). Fractions containing rHMfB were identified by SDS
polyacrylamide gel electrophoresis (SDS-PAGE), and protein in the pooled fractions was
dialyzed against 4x rHMfB binding buffer (40 mM Tris, 4 mM EDTA, 8 mM K2HPO 4,
200 mM NaCl; pH 8.0), and stored at 4' C.
Relaxation ofplasmid DNA. Negatively supercoiled pUC18 plasmid at a
concentration of 50 ýpg/mL was relaxed by treatment with 1 unit of CBE per [Ig plasmid
in 200 mM NaC1, 20 mM Tris, 0.25 mM EDTA and 5% glycerol (pH 8.0) for 1.5 h. at 37
TC. Reactions were stopped by addition of SDS to 1% and proteinase K to 150 [tg/mL
with further incubation at 37 TC for 1.5 h. DNA was further purified by
phenol/chloroform/isoamyl alcohol extraction followed by extraction with chloroform
alone, and then ethanol precipitation. After resuspension in TE buffer, the DNA was
desalted by passage over a NAP-25 column (GE Healthcare), and quantified
spectrophotometrically at 260 nm.
PreparationofsupercoiledplasmidDNA. Prior to large-scale supercoiling
reactions, the optimal ratio of rHMfB to DNA was determined using small-scale test
reactions. For these, 1 ýtg purified relaxed pUC18 at 25 [tg/mL was bound to rHMfB at
ratios between 0.5 and 2.5 [tg protein per [tg plasmid. Binding was carried out for 45
minutes at 370 C in lx rHMfB reaction buffer. After binding, unrestrained compensatory
supercoiling was removed by addition of 10 units of CBE and adjustment of the buffer
with 1/10 volume 590 mM Tris, 8 mM K2HPO 4 , 22 mM EDTA, 100 mM NaCl; pH 8.0
and 1/10 volume H20. Reactions were incubated for a further 1.5 h. at 37 TC. DNA was
then purified using the proteinase K/SDS treatment, phenol/chloroform/isoamyl alcohol
extraction and ethanol precipitation procedures described above. After resuspension,
samples were resolved on a 1% agarose in lx TBE gel. The ratio of protein to DNA that
produced optimal supercoiling was identified, and used for the large-scale reaction.
Conditions in the large-scale reaction were identical to those in the test reactions, with
adjustment of the binding buffer to a final concentration of 350 mM potassium glutamate
for induction of negative supercoils. Unrestrained compensatory supercoiling was
removed by addition of 10 units of CBE and adjustment of the buffer with 1/10 volume
of the adjustment buffer described above, and 1/10 volume of either H2 0 (for positive
supercoils) or 1/10 volume of 1.4 M potassium glutamate (for negative supercoils).
Reactions were purified as before.
Acid phenol extraction to remove nickedplasmid DNA. To remove nicked and
linear plasmid species from the supercoiling preparations, a modified version of the
method of Zasloff et al. (19) was employed. After resuspending the newly supercoiled
plasmid in 1 mM Tris, 1 mM EDTA; pH 8.0, the preparation was desalted by passage
over a NAP-25 column. The DNA solution (<0.5 ýpg/p.L) was adjusted to 75 mM NaCl
and 50 mM sodium acetate (pH 4.0). Molecular biology grade phenol, equilibrated with
50 mM sodium acetate (pH 4.0), was added in equal volume to the salt- and pH-adjusted
DNA solution, and the resulting extraction mixtures were vortexed for 5 min. The
extractions were centrifuged at 6,000 x g for 5 min, and the aqueous layer was
immediately removed and pH-neutralized by adjustment to 200 mM Tris (pH 8.0).
Residual phenol was removed by choloroform extraction, and plasmid was precipitated
with one volume of isopropanol, followed by resuspension in TE buffer.
Analysis ofplasmid topology and topoisomer content in supercoiledsubstrates.
After purification, topoisomer content and direction of supercoiling in the substrates was
determined by 1- and 2-dimensional gel electrophoresis as described elsewhere (20). The
fraction of nicked molecules was determined by gel electrophoresis in the presence of
ethidium bromide.
RESULTS AND DISCUSSION
This method for introducing defined states of supercoiling into covalently closed
circular DNA exploits two phenomena. The first is the constraint of toroidal supercoils
by tetramers of the archaeal histone rHMfB bound to DNA (20). The addition of rHMfB
to closed circular DNA molecule introduces constrained supercoiling of one sign that
must be balanced by induction of an equivalent number of supercoils of the opposite sign
in the unconstrained DNA. The unconstrained compensatory supercoils can be removed
using a DNA topoisomerase, so that the toroidal supercoils constrained by rHMfB are
released as active supercoiling in the closed circular DNA molecules upon removal of the
rHMfB protein molecules (Fig. 2.1). This lab previously made use of this behavior to
develop a method for introducing positive supercoiling into plasmid DNA (15).
I have now exploited a second phenomenon to develop a method to introduce
either positive or negative supercoiling into any closed circular DNA molecule. With
rHMfB, the handedness of the toroids bound by the histone complex is modulated by the
salt composition of the buffer. At low salt concentrations, rHMfB constrains positive
toroids, while higher concentrations (>300 mM K) cause the toroids to reverse in
direction around the rHMfB clusters to constrain negative supercoiling (16). This change
has been attributed to alterations in tetramerization of the protein at the histone fold that
can occur under conditions of high ionic strength (21). Practically speaking, the
dependence of the direction of supercoiling on ionic strength provides a convenient way
to control the sign of supercoiling introduced when generating supercoiled molecules.
The ability of rHMfB to bind DNA with high affinity in either circumstance means that
highly supercoiled plasmid of either sign can be readily produced.
A central feature of this method for preparing supercoiled DNA substrates is the
strict dependence on the use of potassium glutamate as the salt species for controlling the
sign of the supercoiling. We tested several different salt species for efficacy in this
protocol. Of those tested, only potassium glutamate resulted in consistently highly
negatively supercoiled molecules. It is thus recommended for use in preparing negatively
supercoiled substrates. The lack of production of high degrees of negative supercoiling
with other salt species was assumed to be primarily due to inhibition of topoisomerase
activity by the salts in question. To test this hypothesis, we reacted negatively
supercoiled plasmid with increasing amounts of CBE in the absence of rHMfB, using
buffer conditions identical to those present during the relaxation of compensatory
supercoils. Some degree of topoisomerase inhibition exists even without a salt
supplement; this is expected, due to the change of buffer conditions from those which are
ideal for topoisomerase activity to those required for optimal rHMfB binding. Without
exception, the salts that did not produce negatively supercoiled plasmid showed a near
complete inhibition of relaxation of supercoiled plasmid by CBE, even at 10 U CBE / 1
[rg DNA (data not shown).
There are several parameters critical to achieving optimal DNA supercoiling.
One important consideration is that there is an optimal ratio of a protein to DNA that
provides maximal supercoiling of either sign (Fig. 2.2a). At low relative protein
concentrations (<0.8 mass units protein per mass unit of DNA), there are fewer supercoils
constrained by the protein and, as a result, the final product will have a lower average
superhelical density. This importance of this issue is illustrated by the observation at low
salt concentrations that favor positive supercoiling that low levels of negative
supercoiling are actually introduced, rather than the expected positive (20). In the case of
excessive quantities of protein relative to DNA, there is again a reduction in the average
superhelical density introduced into the DNA, presumably as a result of interference with
access of topoisomerases to DNA and thus the ability of the topoisomerase to relax the
unconstrained compensatory supercoiling (20,22). When this optimal ratio is used, large
(>500 [tg) amounts of supercoiled substrate can be produced with a high degree of
superhelical density, with >50% of the DNA molecules in having o-values of 10.03-0.061
(Fig. 2.2b-c).
To achieve this optimal ratio, which is not a threshold ratio, we routinely perform
small-scale (1 [tg of DNA) test reactions by varying the input quantity of rHMfB.
Occasionally, we have encountered circumstances in which the optimal rHMfB:DNA
ratio does not produce maximal supercoiling. In these cases, we have generally found
that further purification of the DNA or an increase in the amount of CBE used is
sufficient to overcome the problem.
A second important parameter involves the amount of CBE for optimal
supercoiling. While a single unit of CBE is sufficient to completely relax 1 [tg of
negatively supercoiled plasmid DNA under standard reaction conditions, significantly
more (-10-fold) is necessary to fully relax the compensatory supercoiling resulting from
rHMfB binding. We attribute this to a combination of factors, including a partial
inhibition of topoisomerase activity under rHMfB binding conditions and a limitation on
the accessibility of the unconstrained DNA to topoisomerase molecules. It is especially
important to prepare the CBE within weeks of the supercoiling procedure to avoid loss of
activity with prolonged storage (months) at -80 oC. Under no circumstances should the
CBE be subjected to freeze-thaw conditions.
While this method for preparing DNA with positive and negative supercoiling
generally results in a population of plasmid molecules with less than 10% nicked form,
we have found that the application of an acid phenol extraction step (19) allows the
preparation of plasmid samples with less than 5% nicked species. We have modified the
acid phenol method of Zasloff et al. by limiting the length of time that the DNA is
exposed to the acidic conditions that can induce depurination. As shown in Figure 2.3, 5
minutes of acid phenol extraction reduce depurination to undetectable levels. This
technique is effective regardless of the superhelical state of the closed-circular molecules
(relaxed, positively or negatively supercoiled), and one extraction step is generally
sufficient to reduce the total fraction of nicked plasmid molecules in the preparation to
<5%.
CONCLUSIONS
We have described a new method for the preparation of large quantities of DNA
substrates containing high levels of either positive or negative supercoiling. The use of
the archaeal histone rHMfB in such a protocol is advantageous, due to its relatively easy
preparation, and ability to reverse binding orientations of DNA in different salt
conditions. Substrates can be prepared via this method that are both highly supercoiled
and free from non-closed circular DNA contaminants, making it ideal for preparation of
substrates for biochemical or biophysical studies.
A
B
C
+
+
1+
I/
~
J
W
Figure 2.1. Reaction scheme for preparation of supercoiled plasmid substrates. A) Relaxed,
closed-circular plasmid is bound with rHMfB in the absence (top) or presence (bottom) of 350
mM potassium glutamate. Toroidal supercoils of one direction are constrained, while
compensatory supercoiling of the opposite sign is unconstrained. B) Topoisomerasecontaining CBE is added without (top) or with (bottom) an additional salt supplement,
removing unconstrained supercoils, while leaving the protein bound supercoils are intact. C)
Bound proteins are removed from the DNA, allowing induced positive (top) or negative
(bottom) supercoils to diffuse throughout the molecules.
rHMfB
B
2in~domns-ro
-
C
-
2 cnseor
2d
EW
-*
Figure 2.2. Optimization and analysis of prepared supercoiled substrates. A) Small-scale
optimization of rHMfB:DNA ratio. At lower levels of rHMfB, low levels of binding occur,
resulting in products with low levels of supercoiling. Above the optimum ratio (denoted with
an *) rHMfB binding interferes with topoisomerase relaxation of compensatory supercoils,
and supercoiling in the product decreases. B) 1- and 2-dimensional analysis of a positively
supercoiled plasmid. In the first dimension, no intercalator is present, and plasmids resolve
on the basis of superhelical density; more supercoiled plasmids travel farther on the gel. In
the second dimension, a low level of intercalator is added, which induces a low level of
positive writhe in the plasmids. While this allows positively supercoiled molecules to travel
farther, negatively supercoiled molecules will have supercoils cancelled out, and thus have
movement retarded. C) 1- and 2- dimensional analysis of negatively supercoiled plasmid,
generated in the presence of potassium glutamate.
preextraction
A
postextraction
OC
CC
untreated
putrescine
D
OC
CC
Figure 2.3. Purification of closed-circular plasmid preparation by extraction with acidic
phenol. A) Damaged plasmid sample before and after acidic phenol extraction (OC = open
circular, CC = closed circular). Extraction reduces the open circular population to less than
5% of the total plasmid. B) Treatment of acidic phenol-extracted plasmid with putrescine.
Incubating extracted plasmid with 0.1 M putrescine (pH 7.0) for one hour at 37 'C does not
introduce any appreciable nicks to the plasmid. Since putrescine converts abasic sites to
strand breaks, this confirms that no significant depurination is occurring during our acidic
phenol extraction.
LITERATURE CITATIONS
1.
Wu, H.Y., Shyy, S.H., Wang, J.C. and Liu, L.F. (1988) Transcription generates
positively and negatively supercoiled domains in the template. Cell, 53, 433-440.
2.
Sinden, R.R., Carlson, J.O. and Pettijohn, D.E. (1980) Torsional tension in the
DNA double helix measured with trimethylpsoralen in living E. coli cells:
analogous measurements in insect and human cells. Cell, 21, 773-783.
3.
Ljungman, M. and Hanawalt, P.C. (1992) Localized torsional tension in the DNA
of human cells. ProcNatl Acad Sci US A, 89, 6055-6059.
4.
Kramer, P.R. and Sinden, R.R. (1997) Measurement of unrestrained negative
supercoiling and topological domain size in living human cells. Biochemistry, 36,
3151-3158.
5.
Lim, H.M., Lewis, D.E., Lee, H.J., Liu, M. and Adhya, S. (2003) Effect of
varying the supercoiling of DNA on transcription and its regulation. Biochemistry,
42, 10718-10725.
6.
Funnell, B.E., Baker, T.A. and Kornberg, A. (1986) Complete enzymatic
replication of plasmids containing the origin of the Escherichia coli chromosome.
JBiol Chem, 261, 5616-5624.
7.
Wang, J.C. (1974) Interactions between twisted DNAs and enzymes: the effects
of superhelical turns. JMol Biol, 87, 797-816.
8.
Bae, S.H., Yun, S.H., Sun, D., Lim, H.M. and Choi, B.S. (2006) Structural and
dynamic basis of a supercoiling-responsive DNA element. Nucleic Acids Res, 34,
254-261.
9.
Bernstein, L.B., Mount, S.M. and Weiner, A.M. (1983) Pseudogenes for human
small nuclear RNA U3 appear to arise by integration of self-primed reverse
transcripts of the RNA into new chromosomal sites. Cell, 32, 461-472.
10.
Levchenko, V., Jackson, B. and Jackson, V. (2005) Histone release during
transcription: displacement of the two H2A-H2B dimers in the nucleosome is
dependent on different levels of transcription-induced positive stress.
Biochemistry, 44, 5357-5372.
11.
Lockshon, D. and Morris, D.R. (1983) Positively supercoiled plasmid DNA is
produced by treatment of Escherichia coli with DNA gyrase inhibitors. Nucleic
Acids Res, 11, 2999-3017.
12.
Gellert, M., Mizuuchi, K., O'Dea, M.H. and Nash, H.A. (1976) DNA gyrase: an
enzyme that introduces superhelical turns into DNA. ProcNatl Acad Sci USA,
73, 3872-3876.
13.
Forterre, P., Mirambeau, G., Jaxel, C., Nadal, M. and Duguet, M. (1985) High
positive supercoiling in vitro catalyzed by an ATP and polyethylene glycolstimulated topoisomerase from Sulfolobus acidocaldarius. Embo J, 4, 2123-2128.
14.
Richardson, S.M., Boles, T.C. and Cozzarelli, N.R. (1988) The helical repeat of
underwound DNA in solution. Nucleic Acids Res, 16, 6607-6616.
15.
LaMarr, W.A., Sandman, K.M., Reeve, J.N. and Dedon, P.C. (1997) Large scale
preparation of positively supercoiled DNA using the archaeal histone HMf.
Nucleic Acids Res, 25, 1660-1661.
16.
Musgrave, D., Forterre, P. and Slesarev, A. (2000) Negative constrained DNA
supercoiling in archaeal nucleosomes. Mol Microbiol, 35, 341-349.
17.
Camerini-Otero, R.D. and Felsenfeld, G. (1977) Supercoiling energy and
nucleosome formation: the role of the arginine-rich histone kernel. Nucleic Acids
Res, 4, 1159-1181.
18.
Starich, M.R., Sandman, K., Reeve, J.N. and Summers, M.F. (1996) NMR
structure of HMfB from the hyperthermophile, Methanothermus fervidus,
confirms that this archaeal protein is a histone. J Mol Biol, 255, 187-203.
19.
Zasloff, M., Ginder, G.D. and Felsenfeld, G. (1978) A new method for the
purification and identification of covalently closed circular DNA molcules.
Nucleic Acids Res, 5, 1139-1152.
20.
Musgrave, D.R., Sandman, K.M. and Reeve, J.N. (1991) DNA binding by the
archaeal histone HMf results in positive supercoiling. Proc Natl Acad Sci US A,
88, 10397-10401.
21.
Marc, F., Sandman, K., Lurz, R. and Reeve, J.N. (2002) Archaeal histone
tetramerization determines DNA affinity and the direction of DNA supercoiling. J
Biol Chem, 277, 30879-30886.
22.
Broyles, S.S. and Pettijohn, D.E. (1986) Interaction of the Escherichia coli HU
protein with DNA. Evidence for formation of nucleosome-like structures with
altered DNA helical pitch. JMol Biol, 187, 47-60.
Chapter 3 - Reactivity of Supercoiled DNA with Chemical and
Enzymatic Probes of DNA Structure
INTRODUCTION
The studies presented in this chapter address biochemical approaches to defining
the structure of positively supercoiled DNA. As discussed in detail in Chapter 1, DNA
supercoiling is ubiquitous in both prokaryotic and eukaryotic cells (1-4), and has been
implicated in a number of important physiological processes, including replication,
transcription, and recombination (5-7). Understanding the structural changes that occur
with the introduction of supercoiling is key to elucidating the mechanisms by which these
physiological effects are exerted. The twin domain model of supercoiling predicts the
presence of both positively and negatively supercoiled DNA in cells (8), but the ready
availability of negatively supercoiled substrate has allowed for a more thorough analysis
than for the positive form.
Negative supercoiling has been shown to produce a number of changes in the
structure of DNA, including localized denaturation and sequence-dependent formation of
alternative secondary structures such as cruciforms, Z-DNA, and triplex DNA (9-12).
However, structural changes that result from positive supercoiling have not been well
defined. Previous studies have suggested that, under certain positive supercoiling
conditions, the bases have an increased solvent exposure (13,14). It has been shown, for
example, that introduction of very high levels of twist using a micromanipulator causes
the DNA to adopt a structure in which a portion of the bases become extrahelical (15).
Findings are limited, however, both in scope and by the methods used to generate
positive supercoiling. Additional studies, with conditions more closely resembling those
in cells, are necessary to better understand the effects of positive supercoiling on the
structure of DNA.
Preliminary studies from this laboratory indicate that the methylating agent
dimethyl sulfate (DMS) is more reactive with positively than negatively supercoiled
plasmid at both the N 7 position of guanine and the N3 position of adenine (16,17). The 3methyladenine result is particularly revealing, since in B-form DNA the N3 position is
buried in the minor groove. When the strands of DNA are more tightly intertwined, as
they are in positive supercoiling, the N3 position could be protected from dimethyl sulfate
alkylation. An increase in methylation at this site could indicate an extrahelical
positioning of at least some of the bases in positively supercoiled DNA.
A number of agents have been identified as probes to elucidate non-canonical
DNA structures. These include single-strand specific nucleases and chemical agents that
react specifically with unpaired bases. S1 nuclease and nuclease BAL-31 are both
capable of cleaving duplex DNA at sites of local melting, including perturbations in base
pairing occurring at a single site (18-20). In addition, the alkylating agent
chloroacetaldehyde has been used to identify non-canonical DNA structures such as the
loop region of cruciforms (21) and the junctions between B- and Z-form DNA (22).
Using these agents along with carefully prepared supercoiled substrates, we have
investigated the effects of positive supercoiling on the structure of DNA. Most
noteworthy, positively supercoiled plasmid is not sensitive to cleavage by single stranded
nucleases, in contrast to the very sensitive nature of negatively supercoiled DNA.
Unfortunately, chloroacetaldehyde results were inconclusive, suggesting that this agent
may be better suited for identifying specific structural phenomena occurring in the DNA.
MATERIALS AND METHODS
Preparationofsupercoiledplasmidsubstrates. Positively supercoiled pUC18
substrates were prepared as described in Chapter 2. Relaxed and negatively supercoiled
substrates were prepared in the same manner as the positive, substituting heat-inactivated
chicken blood extract (CBE) for live enzyme at the appropriate steps to obtain substrates
with the desired presence and direction of supercoiling. All substrates were dialyzed
once in 2 L of TE buffer with 3 M NaC1, using 100,000 MWCO dialysis tubing, followed
by four changes of 2 L of TE buffer, prior to use in chemical or enzymatic experiments.
Correctionfactorsfor quantifying supercoiledsubstratesusing ethidium
fluorescence. To establish correction factors to account for supercoiling-dependent
differences in the binding of ethidium bromide to the plasmid DNA, a gel assay utilizing
both linearized and supercoiled plasmid was performed. For these, a solution of 25 [M
DNA in lx NEbuffer #2 (50 mM NaCl, 10 mM Tris-HC1, 10 mM MgCl 2, 1 mM DTT,
pH 7.9, 100 tig/mL BSA) was prepared, and split in half. To one half, 50 U/mL of
BamHI (New England Biolabs, Ipswich, MA) was added, to the other, an equal volume
of water; this was followed by reaction for 1 hr at 37 °C. Aliquots of 10 [L were
removed from both the linearized and untreated samples, and mixed with 5 tL of
nuclease stop solution. Samples of each intact form were carefully and completely
loaded in adjacent lanes to each of the linearized samples on a 1% agarose in lx TBE gel
containing 0.5 Vig/mL ethidium bromide. Electrophoresis was then performed at 4.5 V/cm
for 90 min. Fluorescence from each band was quantified on an Alpha Innotech
Fluorchem 8900 Imager (San Leandro, CA), and the amount of fluorescence of the intact
form of each plasmid relative to the linearized form was determined.
Nuclease cleavage assays. Reactivity of plasmid DNA with S1 nuclease
(Fermentas, Hanover, MD) was assayed both kinetically and as a dose response. For the
kinetics experiments, 25 [M DNA was reacted with 100 U/mL S nuclease at 37 'C in
lx S1 buffer (40 mM sodium acetate, pH 4.5, 300 mM NaCI, 2mM ZnSO 4). At defined
time points, 10 [tL aliquots were removed and mixed with 10 [tL of nuclease stop
solution (100 mM EDTA, 100 mM Tris-HC1, pH 8.0, 40% sucrose, 0.1% bromophenol
blue) to quench the reaction. Samples were then resolved and quantified on an ethidium
bromide-containing agarose gel. For the dose response, 175 ng of plasmid was reacted
with the nuclease in amounts ranging from 0 to 200 U for 1 hr at 37 "C in lx S1 buffer in
a total volume of 20 [tL. Cleavage was stopped and DNA was quantified as in the
kinetics experiments. To examine the effects of low salt on the reactivity with
supercoiled plasmid, the dose response experiments were repeated with 0-5 U S1
nuclease in lx low salt S1 buffer (40 mM sodium acetate, pH 4.5, 50 mM NaCi, 2 mM
ZnSO 4).
To measure reactivity of supercoiled plasmid with nuclease BAL-31 (New
England Biolabs, Ipswich, MA), the enzyme was reacted at a concentration of 12.5 U/mL
with 25 [tM plasmid at 30 OC in lx BAL-31 reaction buffer (600 mM NaCI, 12 mM
CaC12 , 12 mM MgC12, 20 mM Tris-HC1, 1 mM EDTA, pH 8.0). Reaction was initiated
by addition of the nuclease, and at defined time points 10 [tL aliquots were removed and
the cleavage reaction stopped by addition of 10 1tL of phenol/chloroform/isoamyl
alcohol. After extraction of the aqueous phase, samples were precipitated with an equal
volume isopropyl alcohol, resuspended in TE, and resolved on an ethidium bromidecontaining agarose gel. Relative populations of the different forms of DNA were
quantified using ethidium bromide fluorescence as described above.
Assaying reactivity of chloroacetaldehydewith plasmid substrates. Formation of
etheno adducts on the DNA bases was assayed by reacting 2 ýtg plasmid DNA with
chloroacetaldehyde (Aldrich, St. Louis, MO) at concentrations ranging from 0 to 4.5 mM
in a total volume of 40 [tL of TE buffer for 16 hr at 370 C. Reactions were quenched and
chloroacetaldehyde was removed by precipitation with 2.5 volumes of 100% ethanol,
followed by a 70% ethanol rinse. DNA was resuspended in ddH2 0, and an aliquot was
removed for spectrophotometric quantitation.
The remainder of the DNA was spiked with internal standards consisting of 100
fmol each of 15N-labeled 1,N6-ethenoadenine (EA), and 3,N4 -ethenocytosine (EC)
nucleoside, and digested to deoxynucleoside monophosphates by adjustment to a volume
of 200 tL with sodium acetate buffer (30 mM, pH 5.6, 0.2 mM zinc chloride) and
treatment with 4 units of nuclease P1 at 37 TC for 3 hr. Further digestion to nucleosides
was accomplished by addition of 200 [tL of 30 mM sodium acetate (pH 8.1), 20 units of
alkaline phosphatase, and 1 unit of phosphodiesterase, followed by incubation at 370 C for
6 hr. Enzymes were removed by passage of the reaction mixture over a YM- 10
centrifugal concentrator (Millipore, Billerica, MA), and adducts were isolated and
concentrated using an Agilent 1100 HPLC (Santa Clara, CA).
Analysis was performed on an Agilent model 1100 HPLC equipped with a 1040A
diode array detector coupled to an API 3000 tandem mass spectrometer (Applied
Biosystems, Foster City, CA). All reversed-phase HPLC was carried out on a
Phenomenex C18 column that was eluted isocratically with 98:2 permanganate-treated
H20 (0.1% acetic acid):acetonitrile (0.1% acetic acid). The mass spectrometer was
operated in positive ion mode; the first quadropole was set to transmit the precursor ions
MH+ at m/z 255 and 281 for the 15N-EC and eA internal standards, and at m/z 252 and
276 for the EC and EA analytes. The product ions were monitored in the third quadropole
at m/z 139 and 165 for the 15N-labeled internal standards, and 136 and 160 for the
analytes.
RESULTS
Correctionfactorsfor supercoiledplasmid. The binding of ethidium to DNA
causes a localized unwinding at the site of intercalation. For linear DNA, the
compensatory overwinding in the remaining DNA that occurs as a result of ethidium
binding can diffuse out of the ends of the molecule, so the overall binding is limited only
by the number of available binding sites on the DNA. A covalently closed-circular
molecule presents a different situation. In this case, the global overwinding that
compensates for the localized unwinding must remain in the molecule, the energetics of
which effectively translate into a positive writhe. There is a maximal amount of positive
writhe that can be introduced into a closed-circular molecule, which limits the binding of
ethidium to the plasmid molecule (23). As a result, not only will closed-circular DNA
bind ethidium differently than an equivalent linear molecule, but the resultant supercoiled
states will also vary. Positively supercoiled plasmid will be able to bind the least amount
of ethidium before reaching a point where the molecule cannot accommodate any more
introduced supercoils. Negatively supercoiled plasmid will bind considerably more
ethidium, although not as much as an equivalent linear molecule; relaxed will bind an
intermediate amount.
Since measurements based on ethidium fluorescence are dependent on the amount
of bound ethidium, it is crucial to account for this differential binding to obtain accurate
quantification. To ensure that measurements were accurate for quantifying the differently
supercoiled molecules, ethidium fluorescence of equal amounts (determined
spectrophotometrically) of the three linearized forms of plasmid on a gel were compared
at saturating ethidium concentrations. The quantified bands were virtually identical, with
variations small enough to be accounted for by pipetting errors. Having established that
A 260
readings were sufficient to quantify DNA and that the same amount of plasmid was
present in each sample, fluorescence of the relaxed, positively and negatively supercoiled
plasmid were compared. As expected, negatively supercoiled plasmid was the most
fluorescent, followed by relaxed, and then positively supercoiled DNA. The relative
fluorescence (compared to negatively supercoiled plasmid) of relaxed and positively
supercoiled samples was 0.67 and 0.5, respectively. Previous studies determined that
linearized pUC 19 is 1.4 times more fluorescent than the natively negatively supercoiled
molecule (24). This is in general agreement with our results, defining the correction
factors for closed circular molecules as follows: negatively supercoiled plasmid: 1.40;
relaxed plasmid: 2.09; positively supercoiled plasmid 2.54.
Positively supercoiledplasmidis not cleaved at an increasedrate by single strand
specific nucleases. S nuclease is a glycoprotein from Aspergillus oryzae that has both
exo- and endonuclease activity. It exhibits endonuclease activity at regions of singlestranded DNA, to the level of a single unpaired base (25). In all conditions tested,
negatively supercoiled plasmid was highly reactive with S1 nuclease, presumably due to
the underwinding and associated transient denaturation of the DNA strands (Figs. 3.1-3).
This is consistent with published reports (20,25,26). In the kinetics experiment, both
relaxed and positively supercoiled plasmid exhibited significantly less reactivity than
negative, with relaxed plasmid reacting slightly more efficiently than the positively
supercoiled form (Fig. 3.1). In addition, the reactivity of relaxed and positive is similar
in the dose response experiments (Fig. 3.2), which may possibly result from cooperative
enzyme activity at higher concentrations rather than any single-stranded character of the
DNA. Reducing the ionic strength increased the reactivity of all three substrates
considerably (data not shown), consistent with previous studies that found higher levels
of cleavage at low ionic strength (20). While the overall reactivity of all topological
forms of DNA increased with lower ionic strength, no differences between positively
supercoiled and relaxed plasmid emerged. Lowering the ionic strength reduces the
shielding of the highly charged phosphate backbone by counterions, which prevents the
segments in the helix from coming into as close proximity as occurs at higher ionic
strength. As a result, the writhe component in the molecule is reduced and as the twist
must increase to compensate (27). The lack of a difference in reactivity between relaxed
and positive substrates at lower ionic strength suggests that even though the writhe is
decreasing at lower salt, the resulting increase in twist is not responsible for the increased
reactivity, as the relaxed molecules show an increase in cleavage equivalent to that seen
in positive.
Nuclease BAL-31 from Alteromonas espejianaalso possesses both exo- and
endonuclease activity, but it is active at a pH that is much closer to neutrality than S (pH
8.0 vs. pH 4.5) (19). Again, the negatively supercoiled plasmid was the most reactive,
with relaxed and positively supercoiled plasmid being cleaved at significantly slower
rates (Fig. 3.3). At this low enzyme concentration, the relaxed plasmid was cleaved
slightly more efficiently than positive, much as it was in the S1 nuclease kinetics
experiments.
Neither S1 nor BAL-31 showed an increase in reactivity with positively
supercoiled plasmid, relative to the relaxed form. In fact, at low concentrations of either
enzyme, such as were used in the time course studies, positively supercoiled substrates
were cleaved at a slower rate than the relaxed form. While these findings are not
consistent with extrahelical bases in positively supercoiled substrates, it is possible that
structural changes induced by positive supercoiling interfere with the binding or activity
of these enzymes. Although both enzymes have been shown to cleave at sites of unpaired
bases (13,25), it is not clear exactly what the enzymes recognize in terms of DNA
structure. A structure in which extrahelical bases are present in positively supercoiled
DNA may be sufficiently different from the structure being recognized in negatively
supercoiled plasmid that it would not be cleaved by either enzyme, particularly if the
backbone was shielded at the center of the helix.
Chloroacetaldehyde is not an effective probe for transientsingle-strandedregions
in DNA. Chloroacetaldehyde is a vinyl chloride metabolite that reacts with the hydrogen-
bonding region of bases to form four different exocyclic etheno adducts (Fig. 3.4)
(28,29). The formation of both 1,N6-ethenodeoxyadenosine and 3,N4-ethenocytosine
were analyzed using LC-MS (Fig. 3.5). While there appears to be a slight increase in the
quantities of both adducts in the negatively supercoiled plasmid, this effect is not
statistically significant and it is not as pronounced as would be expected based on the
single-strand nuclease data. This may be due to the multi-step reaction mechanism
proposed for the formation of the etheno adducts. The details of the reaction are still in
dispute, but a generally accepted scenario involves attack of the chloroacetaldehyde at the
exocyclic amino group on the base to form an unstable intermediate, which then reacts
with a heterocyclic nitrogen to form a cyclic adduct that finally undergoes a dehydration
step to form an etheno adduct (30,31). Even if the molecule were able to undergo an
initial attack, the nature of the solvent exposure of the base could be incompatible with
subsequent cyclization and dehydration steps. Because this entire process is necessary to
form the analytes detected here, chloroacetaldehyde may not be an ideal agent to measure
base solvent exposure.
Chloroacetaldehyde has been used to identify sites of single-stranded character in
several instances (21,22), but this is generally as a qualitative agent. Typically,
chloroacetaldehyde-reacted DNA is linearized, and then treated with S1 nuclease to
express the adducted base as a strand break (21), although Maxam-Gilbert sequencing
methods are used, as well (22). Such methods are useful for identifying specific sites that
are susceptible to damage, such as the loop region of cruciforms, but the presence of
multiple adduction sites throughout the DNA would complicate interpretation and
prevent a good quantitative assessment of base unpairing. Interestingly, the most
informative aspect of chloroacetaldehyde reactivity may be in the partitioning between
the two guanine adducts. After the initial attack on the exocyclic nitrogen, the adduct can
cyclize with either of two ring nitrogens, the N' and the N3 . Increased hydrogen bonding
will interfere with cyclization at the N' position, shifting the reaction to favor cyclization
at the N3 (32). Measuring changes in the ratios between these two adducts may provide
key information about the hydrogen bonding state of guanine in supercoiled DNA.
It is worth noting that a higher dose of chloroacetaldehyde was tested than is
shown in Figure 3.5 (data not shown). In all cases, the dose-response deviated from
linearity at this higher dose, showing a substantial increase in the rate of formation of
adducts. This could be due to a "seeding" effect due to the inability of an ethenoadducted base to return to a normal base-pairing geometry. This perturbation may serve
to destabilize neighboring base pairs, making those bases more solvent exposed and thus
more susceptible to chloroacetaldehyde attack. Mismatches involving canonical base
pairs are known to destabilize the helix (33,34); so it is reasonable to expect bulky etheno
adducts to have a similar, if not more pronounced, effect.
DISCUSSION
Biomechanical and biochemical experiments have suggested that sufficient levels
of positive supercoiling can result in DNA structures in which the bases are positioned
extrahelically. We have investigated the possibility that such structures may occur at
physiologically relevant levels of positively supercoiling by reacting supercoiled and
relaxed plasmid substrates with enzymatic and chemical agents that are selective for
unpaired bases and single stranded structure.
No evidence was found in these studies for extensive or obvious solvent exposure
of bases induced by the levels of positive DNA supercoiling that are introduced with our
method (c = +0.04). There are several possible explanations for this result. The simplest
among these is the absence of solvent exposed bases under physiologically relevant
conditions of torsional tension, at least at the superhelical density of the substrates
utilized in these assays (c = +0.04). It could be that the threshold for transition to an
extrahelical base structure occurs at superhelical density values that are too high for any
extrahelical bases to be detected in these substrates. It is also possible that the methods
of analysis were insufficient to detect extrahelical bases that were nonetheless present
transiently, perhaps as a result of fast kinetics of base flipping. Recognition of
overwinding-induced structural changes by single-strand specific nucleases has not been
established, so the possibility of such changes being present cannot be ruled out.
Chloroacetaldehyde was not effective in identifying increased reactivity with negative
supercoiling, where solvent exposed bases are known to occur, so the lack of increased
reactivity of bases in positively supercoiled plasmid is not conclusive in these studies.
Further study with additional chemical probes may clarify what structural changes
occur when DNA is subjected to overwinding. Repeating the DMS studies that were
initially performed in this laboratory (16,17) with our improved substrate, and expanding
them to include relaxed as well as supercoiled plasmid could be informative. A number
of other chemical agents are well established as probes of stably unpaired structure,
including potassium permanganate (35) and diethyl pyrocarbonate (36). Comparing
reactivity of these agents with relaxed and supercoiled plasmids should help broaden the
picture of structural changes introduced with positive supercoiling.
Still, the structural changes that occur when DNA is positively supercoiled may
be too subtle at the level of supercoiling present in these studies to be detected using
chemical or enzymatic means. For this reason, complementary approaches should be
considered that would allow for both detection of subtle structural changes, as well as
introduction of precise and relatively high levels of twist into the DNA. In the following
chapters, these approaches are addressed. Raman spectroscopy is utilized to examine the
subtle structural changes occurring in supercoiled plasmid substrates, giving detailed
information about precisely which bonds are being affected. Preliminary substrate design
and considerations for fluorescence-based micromanipulation experiments are then
discussed, providing a foundation for experiments that will allow for elucidation of
structural changes as a function of introduced twist.
75
* positive
* negative
A relaxed
50
25
,
.'•I1m
7
ýW
'
I
20
40
Time (minutes)
60
Figure 3.1. Kinetics of reaction of S 1 nuclease with supercoiled and relaxed substrates.
Negatively supercoiled plasmid is cleaved by the single-strand specific S nuclease at a much
faster rate than either relaxed or positively supercoiled plasmid. Plasmid substrates were
reacted with S nuclease as described in Materials and Methods. Data points shown are the
average of four replicates; error bars represent the standard error.
100
A-~~-~-
--
I'1~1-
M
75
* positive
* negative
A relaxed
Eb-
50
100
U Si nuclease
150
200
Figure 3.2. Reaction of S1 nuclease with supercoiled and relaxed substrates. Negatively
supercoiled plasmid is cleaved by the single-strand specific S1 nuclease considerably more
than either relaxed or positively supercoiled plasmid. Plasmid substrates were reacted with
S1 nuclease as described in Materials and Methods. Data points shown are the average of
four replicates; error bars represent the standard error.
4 tf%
IUV
-
C
, 75
L,.
Uc
0U
50
Spositive
Snegative
C"
Arelaxed
25
04
0
1
10
20
30
Time (minutes)
Figure 3.3. Kinetics of reaction of nuclease BAL-31 with supercoiled and relaxed substrates.
Negatively supercoiled plasmid is cleaved by the single-strand specific nuclease BAL-31 at a
much faster rate than either relaxed or positively supercoiled plasmid. Plasmid substrates
were reacted with nuclease BAL-31 as described in Materials and Methods. Data points
shown are the average of at least four replicates; error bars represent the standard error of the
data.
NH2
N
NNI
N
NH2
N
N
'N"-
O
N
0
CI
Chloroacetaldehyde
O
N
NH
N
O
N
NH2
N
NH
N N N
Figure 3.4. Reaction of chloroacetaldehyde with DNA bases. Chloroacetaldehyde reacts with
adenine, cytosine, and guanine to form 1, N6 -ethenoadenine, 3,N4 -ethenocytosine, 1,N2ethenoguanine and N2 ,3-ethenoguanine.
12
* positive
negative
A relaxed
ii
ol 0
4
k
0
.1
0.5
mM chloroacetaldehyde
·--
* positive
9 negative
A relaxed
Al r9 IlL
~--------i
I
0
I
·
0.5
1
mM chloroacetaidehyde
Figure 3.5. Reaction of chloroacetaldehyde with supercoiled and relaxed substrates. No
significant effect is seen on the formation of etheno adducts as a function of supercoiling.
Plasmid substrates were treated with chloroacetaldehyde as described in Materials and
Methods. Data points shown are the average of two replicates; error bars represent the
standard error.
LITERATURE CITATIONS
1.
Wu, H.Y., Shyy, S.H., Wang, J.C. and Liu, L.F. (1988) Transcription generates
positively and negatively supercoiled domains in the template. Cell, 53, 433-440.
2.
Figueroa, N. and Bossi, L. (1988) Transcription induces gyration of the DNA
template in Escherichia coli. ProcNatl Acad Sci US A, 85, 9416-9420.
3.
Brill, S.J. and Sternglanz, R. (1988) Transcription-dependent DNA supercoiling
in yeast DNA topoisomerase mutants. Cell, 54, 403-411.
4.
Giaever, G.N. and Wang, J.C. (1988) Supercoiling of intracellular DNA can occur
in eukaryotic cells. Cell, 55, 849-856.
5.
Funnell, B.E., Baker, T.A. and Kornberg, A. (1986) Complete enzymatic
replication of plasmids containing the origin of the Escherichia coli chromosome.
JBiol Chem, 261, 5616-5624.
6.
Steck, T.R., Franco, R.J., Wang, J.Y. and Drlica, K. (1993) Topoisomerase
mutations affect the relative abundance of many Escherichia coli proteins. Mol
Microbiol, 10, 473-481.
7.
Droge, P. (1993) Transcription-driven site-specific DNA recombination in vitro.
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Liu, L.F. and Wang, J.C. (1987) Supercoiling of the DNA template during
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Kowalski, D., Natale, D.A. and Eddy, M.J. (1988) Stable DNA unwinding, not
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Lilley, D.M. (1981) Hairpin-loop formation by inverted repeats in supercoiled
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Peck, L.J., Nordheim, A., Rich, A. and Wang, J.C. (1982) Flipping of cloned
d(pCpG)n.d(pCpG)n DNA sequences from right- to left-handed helical structure
by salt, Co(III), or negative supercoiling. Proc Natl Acad Sci US A, 79, 45604564.
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Beltran, R., Martinez-Balbas, A., Bernues, J., Bowater, R. and Azorin, F. (1993)
Characterization of the zinc-induced structural transition to *H-DNA at a
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Lau, P.P. and Gray, H.B., Jr. (1979) Extracellular nucleases of Alteromonas
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McClellan, J.A. and Lilley, D.M. (1991) Structural alteration in alternating
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Allemand, J.F., Bensimon, D., Lavery, R. and Croquette, V. (1998) Stretched and
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LaMarr, W.A. (1998) Thesis Ph. D. --Massachusetts Institute of Technology
Division of Toxicology 1998.
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LaMarr, W.A. Unpublished observations.
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Shenk, T.E., Rhodes, C., Rigby, P.W. and Berg, P. (1975) Biochemical method
for mapping mutational alterations in DNA with S1 nuclease: the location of
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Gray, H.B., Jr., Ostrander, D.A., Hodnett, J.L., Legerski, R.J. and Robberson,
D.L. (1975) Extracellular nucleases of Pseudomonas BAL 31. I. Characterization
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Wiegand, R.C., Godson, G.N. and Radding, C.M. (1975) Specificity of the S1
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Dayn, A., Malkhosyan, S., Duzhy, D., Lyamichev, V., Panchenko, Y. and Mirkin,
S. (1991) Formation of (dA-dT)n cruciforms in Escherichia coli cells under
different environmental conditions. JBacteriol,173, 2658-2664.
22.
Kohwi-Shigematsu, T., Manes, T. and Kohwi, Y. (1987) Unusual conformational
effect exerted by Z-DNA upon its neighboring sequences. ProcNatl Acad Sci US
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23.
Radloff, R., Bauer, W. and Vinograd, J. (1967) A dye-buoyant-density method for
the detection and isolation of closed circular duplex DNA: the closed circular
DNA in HeLa cells. Proc Natl Acad Sci US A, 57, 1514-1521.
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Milligan, J.R., Aguilera, J.A. and Ward, J.F. (1993) Variation of single-strand
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solution. Radiat Res, 133, 151-157.
25.
Beard, P., Morrow, J.F. and Berg, P. (1973) Cleavage of circular, superhelical
simian virus 40 DNA to a linear duplex by S 1 nuclease. J Virol, 12, 1303-1313.
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Dasgupta, S., Allison, D.P., Snyder, C.E. and Mitra, S. (1977) Base-unpaired
regions in supercoiled replicative form DNA of coliphage M13. JBiol Chem, 252,
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Bednar, J., Furrer, P., Stasiak, A., Dubochet, J., Egelman, E.H. and Bates, A.D.
(1994) The twist, writhe and overall shape of supercoiled DNA change during
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Possible implications for DNA structure in vivo. JMol Biol, 235, 825-847.
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Kochetkov, N.K., Shibaev, V.N. and Kost, A.A. (1971) New reaction of adenine
and cytosine derivatives, potentially useful for nucleic acids modification.
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Sattsangi, P.D., Leonard, N.J. and Frihart, C.R. (1977) 1,N2-ethenoguanine and
N2,3-ethenoguanine. Synthesis and comparison of the electronic spectral
properties of these linear and angular triheterocycles related to the Y bases. J Org
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30.
Sattsangi, P.D., Barrio, J.R. and Leonard, N.J. (1980) 1,N6-Etheno-bridged
adenines and adenosines. Alkyl substitution, fluorescence properties, and
synthetic applications. Journalof the American Chemical Society, 102, 770-774.
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deoxyribopolynucleotides. 1. Reaction products. Biochemistry, 21, 5717-5722.
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Itakura, K. and Breslauer, K.J. (1982) Structure, dynamics, and energetics of
deoxyguanosine. thymidine wobble base pair formation in the selfcomplementary d(CGTGAATTCGCG) duplex in solution. Biochemistry, 21, 437444.
34.
Aboul-ela, F., Koh, D., Tinoco, I., Jr. and Martin, F.H. (1985) Base-base
mismatches. Thermodynamics of double helix formation for dCA3XA3G +
dCT3YT3G (X, Y = A,C,G,T). Nucleic Acids Res, 13, 4811-4824.
35.
Sasse-Dwight, S. and Gralla, J.D. (1989) KMnO4 as a probe for lac promoter
DNA melting and mechanism in vivo. J Biol Chem, 264, 8074-8081.
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Leonard, N.J., McDonald, J.J., Henderson, R.E. and Reichmann, M.E. (1971)
Reaction of diethyl pyrocarbonate with nucleic acid components. Adenosine.
Biochemistry, 10, 3335-3342.
Chapter 4 - Raman Spectroscopic Studies of Positive Supercoiling
INTRODUCTION
Unconstrained supercoiling in cellular DNA is ubiquitous (1-3), resulting from
the forced rotation of chromosomal DNA by anchored, helix-tracking polymerases and
other enzymes (4). Both positive and negative supercoiling are generated during these
processes (5), and both can effect vital cellular processes, including transcription,
replication and recombination, among others (6-9). Establishing the effects of DNA
supercoiling on the structure of the molecule is critical to gaining a more complete
understanding of the resultant physiological effects, since the activity of DNA is so
intimately tied to its structure.
In the previous chapter, chemical and enzymatic probes that are sensitive to
altered DNA secondary structure were used to study structural changes in positively and
negatively supercoiled DNA. While negatively supercoiled DNA showed increased
reactivity to single strand-specific nucleases, no changes were seen with positively
supercoiled DNA. Studies with chemical probes were largely inconclusive. Taken
together, these studies suggest that structural changes that occur with positive
supercoiling in DNA are subtle and may not be identifiable with these relatively crude
structural probes that are usually employed to identify more overt structural changes, like
cruciform and Z-DNA formation.
Because of the potentially subtle nature of the structural changes occurring in
positively supercoiled DNA, a more sensitive method of detection was required. To this
end, spectroscopic and crystallographic methods could provide more detailed
information, but methods such as NMR and X-ray crystallography that are often used to
determine macromolecular structure are not applicable to supercoiled DNA because of
the size and flexibility of supercoiled plasmid molecules. Our current DNA supercoiling
method is limited to fairly large DNA molecules (>500 base pairs), which tumble too
slowly on the NMR time scale and are too complicated for crystallographic methods.
Fortunately, Raman spectroscopy does not suffer from these same limitations
(10). This method has been successfully applied to investigate changes in DNA structure
caused by a multitude of factors, including sequence effects, formation of non-canonical
DNA structures, and the binding of proteins and small molecules to DNA (11-18). DNA
has a well-studied Raman signature: there is a significant body of work on the
contribution of the different vibrational modes to the overall spectrum, which provides
insight concerning specific and subtle changes that occur when a structural effector is
introduced. Since the intensity of a given Raman band is proportional to the population
of that specific molecular species present in the solution being probed, both the identity
and the degree of structural change can be determined by carefully monitoring peak
frequencies and intensities.
In this study, highly purified supercoiled plasmid substrates were prepared and, in
collaboration with colleagues at the University of Missouri at Kansas City, the Raman
spectra of positively and negatively supercoiled plasmids were compared to the spectrum
of the relaxed form. Resulting data show that all supercoiling states of the plasmid are
predominately B-form DNA, with subtle but reproducible changes induced by
supercoiling. Changes are observed primarily in the DNA sugar-phosphate backbone.
MATERIALS AND METHODS
Preparationofsupercoiledand relaxed samples. Positively supercoiled pUC18
plasmid substrates were prepared as described in Chapter 2. Negatively supercoiled and
relaxed plasmids were prepared in the same fashion, substituting heat-inactivated CBE
for active enzyme at the appropriate relaxation steps to arrive at the final desired
supercoiling state. Following purification, all plasmid preparations were dialyzed once
against 2 L of 3 M NaCl in TE (10 mM Tris, pH 8.0, 1 mM EDTA) using 100,000
molecular weight cut-off tubing, followed by 4 buffer changes of 2 L 200 mM NaCi
adjusted to p-I 7.4.
Samples were concentrated to a final concentration of 25-40 mg/mL using
Microcon YM-30 centrifugal concentrators (Millipore, Billerica, MA). Prior to use,
concentrators were washed with 0.1 N NaOH, followed by several rinses with H20 to
removed residual glycerin from the membranes. For deuterated samples, concentrated
plasmids were lyophilized to dryness, and redissolved in D20 (D, 99.9%) (Cambridge
Isotope Laboratories, Inc., Andover, MA) to the pre-lyophilization volume. In both
cases, pH or pD was adjusted prior to preparation of sample cells: pH 7.4 in the case of
hydrated samples, and pD 7.0 in the case of deuterated samples. For the sample cells, 812 tiL of concentrated sample was sealed in a glass capillary tube.
Collection of Raman spectra. Raman spectra were collected on two different
instruments utilizing different excitation wavelengths. Spectra were collected on a Spex
500-M single spectrograph (Metuchen, NJ) equipped with a liquid nitrogen-cooled, back-
thinned, charge-coupled-device detector (Spectrum One, Instruments S.A., Edison, NJ).
Spectra were excited at 532 nm with a solid state Nd:YVO 4 laser (Verdi, Coherent, Santa
Clara, CA). Spectra were also collected using near-infrared excitation on a Kaiser Optics
HoloSpec VPT spectrometer (Ann Arbor, MI) equipped with a liquid nitrogen-cooled,
charge-coupled-device detector optimized for the near-infrared region (Roper, model
1024EHRB, Trenton, NJ). Spectra were excited at 752 nm with a Ti:sapphire laser
(Coherent, Model 890), which was pumped at 532 nm (Spectra-Physics, model Millennia
X, Mountain View, CA). Raman wavenumbers were calibrated using the emission lines
from a neon lamp and the 459 cm- 1 band of liquid CC14 and are accurate to ± 1 cm'1 .
Samples were thermostatted at 17 OC during data collection. Data presented represent an
average of approximately 200 exposures of 30 s each.
Processingof Raman spectral data. Following spectral averaging, weak
scattering by the aqueous solvent (200 mM NaCl in either H2 0 or D20) was removed and
baselines redrawn using established methods (19). The Raman marker of the DNA
phosphate group at 1092 cm - 1 was employed for spectral intensity normalizations, as also
described previously (13).
RESULTS
Considerationsfor sample preparation. Laser Raman spectroscopy is a powerful
technique that is highly sensitive to contamination by non-analyte molecules possessing
covalent bonds. Because of this, the utmost care must be taken in the preparation and
purification of molecules for Raman studies. Two species of particular concern are Tris
cation and glycerol. Tris is commonly used as a buffer in the preparation and
manipulation of nucleic acids. It has a distinctive Raman signature with prominent bands
at 757, 894, 1064 and 1468 cm 1', all of which could potentially interfere with
interpretation of the Raman spectra of the DNA (20). Careful removal of Tris by dialysis
against a high salt buffer followed by repeated buffer changes of the final analysis buffer
(200 mM NaC1) is sufficient to remove the cation and prevent it from interfering with the
Raman signal from the supercoiled target.
Another potential source of contamination is glycerin, which is present in small
amounts in the centrifugal concentrator membranes to allow for dry storage. Washing
concentrator membranes with 0.1 N NaOH followed by repeated rinses with H2 0 is
effective in removing the glycerin and preventing this species from interfering with the
subsequent Raman analyses.
Finally, great care should be taken to avoid contamination with even small
amounts of fluorescent species. The nature of Raman scattering, in which -1 in 107
photons is scattered inelastically, requires an intense excitation source to get a relatively
small signal. Because of this, even slight contamination with fluorescent molecules can
lead to significant background fluorescence in Raman spectra. Such fluorescence can
dramatically reduce the signal to noise ratio. Potential sources of fluorescent
contaminants include oxidation products from phenol used in DNA purification
procedures, and plasticizers present in H20 from some sources. Using fresh reagents for
substrate purification and ultra-pure H2 0 can significantly reduce fluorescent
contamination. The use of near-IR (752 nm) excitation can also dramatically reduce the
background from fluorescent contaminants, as these contaminants are less likely to be
excited at the longer wavelength.
Raman spectra of negatively supercoiledand relaxedplasmid. Raman spectra of
negatively supercoiled and relaxed pUC 18 plasmid and their resultant difference spectra,
obtained at 752 and 532 nm, are displayed in Figures 4.1 and 4.2, respectively. Parent
spectra had minimal fluorescence and high signal-to-noise ratios that permitted
generation of high quality difference spectra.
Raman spectra obtained at both excitation wavelengths reveal that negatively
supercoiled and relaxed plasmids exist predominantly in the B-form. The Raman band at
680 cm-1 , indicative of the C2'endo/anti deoxyribose conformation of guanosine
nucleosides, the 790 and 833 cm' bands that are indicative t/t phosphodiester torsions
in the gauche-/gauche-range, and the C2'H 2 band at 1421 cm-1 are all reliable markers of
B-form DNA (21), and are clearly present in both negatively supercoiled and relaxed
plasmids. The absence of a C3' endo/syn Raman marker band near 625 cm-1 confirms the
absence of Z-DNA in both cases (11), as would be expected based upon the sequence of
the pUC18 plasmid.
The difference spectra indicate that there are subtle but distinct changes in
conformation between negatively supercoiled and relaxed plasmids. These
conformational differences can be attributed primarily to modest changes in DNA
backbone conformation that accompany the introduction of negative torsion, although
some disruption in hydrogen bonding is also apparent. Specific features of the difference
spectra will be discussed in detail in the Discussion.
Raman spectra ofpositively supercoiledplasmid. In Figures 4.3 and 4.4, the
Raman spectrum obtained on the positively supercoiled plasmid is compared to the
spectrum of the relaxed plasmid, at 752 and 532 nm excitation wavelengths, respectively.
As with negatively supercoiled and relaxed samples, the Raman spectrum of the
positively supercoiled plasmid indicates that the B form predominates, with Raman
marker bands evident at 680, 790, 833 and 1421 cm-'. The difference spectra again show
subtle differences between positively supercoiled and relaxed plasmids, although the
differences are not identical to those seen with negative supercoiling. Again, specific
features of the difference spectra will be detailed in the Discussion.
Raman spectra ofsupercoiledand relaxed substratesin D2 0. To confirm the data
obtained with hydrated samples, Raman scattering measurements were repeated with
deuterated samples (Figs 4.5 and 4.6). In all cases the results obtained on hydrated
samples were reinforced, although spectral baselines for the deuterated samples were not
nearly as good as for the hydrated samples. The reason for this is partial hydration of the
deuterated samples due to humidity in the air. Even with minimization of exposure of the
deutrated samples to the environment, humidity in the air can rapidly exchange with the
solvent. In hydrated samples, the contribution of H20 to the spectra can be removed by
subtracting the spectra of 200 mM NaCl in H20 from the sample spectra. Ideally, the
deuterated samples could be corrected for solvent contributions by subtracting a spectrum
of 200 mM NaCl in D2 0. However exchange of solvent deuterons with protons from the
environment results in the solvent contribution being a combination of D2 0 and HDO,
which have distinctly different scattering characteristics. The proportion of D20 to HDO
is not identical from sample to sample, leading to the difficulty in solvent correction and
the uneven spectral baselines seen in Figures 4.5 and 4.6.
DISCUSSION
Generation ofRaman difference spectra. Raman difference spectroscopy is a
highly sensitive technique that can be used to identify subtle and specific differences in
DNA structure with the introduction of an effector of DNA structure. Because of the
sensitivity of this technique, it is crucial to eliminate any extraneous differences in the
substrates being compared to ensure that all differences observed are a result of the
introduction of the parameter being studied. By carefully preparing and purifying the
substrates, and comparing supercoiled to relaxed (rather than linear) plasmid of the same
sequence, we have hopefully eliminated all causes for the observed difference bands,
other than the supercoiled state of the molecule. Since the same difference peaks are
arising at both excitation wavelengths, the appearance of these peaks is a real
phenomenon, and not merely noise. Further confirmation comes from the deuterated
spectra. Negatively supercoiled and relaxed spectra have been obtained in duplicate at
both excitation wavelengths; positively supercoiled plasmid awaits another replicate.
Changes in DNA structure with the introduction ofsupercoiling:Base stacking.
Several bands corresponding to base moieties exhibit hypochromism upon base
unstacking, the most sensitive of which are the 727 cm' band of adenine and the 1236
cm-1 band of thymine (22,23). No significant intensity increase is observed at either of
100
these positions in difference spectra of negatively or positively supercoiled plasmids
(Figures 4.1 and 4.3, respectively), indicating that base stacking is not significantly
disrupted in either case.
Changes in DNA structure with the introductionofsupercoiling: Base pairing.
The peak shift from -1483 to 1495 cm-' in both supercoiled states suggests a decrease in
the hydrogen bonding strength at the guanine N7 position (16,17). Since the GN7 position
is located in the major groove, a change in the hydrogen bonding status at this site may
reflect an increase in exposure of the groove to solvent molecules. Such an increase in
major groove exposure might be expected with the increase in DNA bending and twisting
that accompany supercoiling. Recently, it has been hypothesized that this shift may also
involve a contribution from the hydrogen bonding state of the 06 position in guanine, so
this shift may also suggest a decrease in the stability of G:C base pairing in both
supercoiled states.
Positively supercoiled plasmid also shows a slight band shift from 1574 to 1581
n . The 1577 cm - 1 band in DNA results
cm'
from the contributions of two modes, the N6H2
scissor vibration of adenine, and the adenine ring (24). The scattering from the N6 H2
scissor is very sensitive to deuteration, shifting to 1190 cm-', which is not seen here.
Since the bandshift remains at the 1574 to 1581 cm-' location in the deuterated sample,
the shift is due to the contribution from the purine ring (25). Such a shift is associated
with a disruption of the hydrogen bonding state of the moiety, so this may represent a
decrease in stability of A:T base pairing in positively supercoiled DNA.
Changes in DNA structure with the introductionof supercoiling: Effects on DNA
backbone conformation. Both positively and negatively supercoiled plasmids show an
101
increase in intensity at 792 cm -1', a band associated with ca/ý phosphodiester torsions in
the gauche-lgauche-range (21,25). In pUC18, which has -50% GC content, the
phosphodiester torsion is overlapped by contributions from both cytosine (784 cm 1') and
thymine (790 cm' 1) (26). The intensity change, however, is likely due to perturbation of
the phosphodiester torsion, since the shift is not noticeably sensitive to deuteration, as it
would be if it were due to either of the base moieties.
The peak at 854 cm-1 that appears with negative but not positive supercoiling
indicates a shift in the a/13/y backbone torsions from gauche-/trans/gauchet to a
trans/trans/transconformation for some phosphodiester moieties (27). This band has
been shown previously to be influenced by helical unwinding, which is consistent with its
appearance in negatively but not positively supercoiled DNA.
Positive supercoiling results in an increase in intensity at 680 cm " 1 that is not
observed with negative supercoiling. This peak is diagnostic of C2' endo/anti
deoxyguanosine nucleosides that are characteristic of B-form DNA (11). A
corresponding trough at 811 cm -~ induced by positive supercoiling is also consistent with
a small reduction in the number of sugar moieties in a C3' endo/anti conformation, a
conformation that is generally associated with A-form DNA (28). Additionally, both
positive and negative supercoiling result in an increase in intensity in the 1340-1345 cm - 1
range, with the peak exhibiting greater intensity with positive supercoiling. The 1345
cmr1 band is associated with C2'endo/anti sugar pucker in deoxyadenosine (dA)
nucleosides (25,28). Taken together, the data suggest that a small number of nucleosides
with A-form character are converting to the B-form with the introduction of positive
supercoiling. The presence of C3' endo/anti deoxynucleosides in relaxed mixed-
102
sequence B DNA is not unprecedented. Although the predominant nucleoside
conformation is expected to be C2' endo/anti for B DNA, a fraction of the furanose rings
may exist in a. C3' end/anti conformation (29). The conversion of these moieties to a C2'
endo/anti conformation as seen here is consistent with the anticipated lengthening of the
backbone that is required when DNA accommodates positive supercoiling. When DNA
is overwound, the sugar-phosphate backbone must lengthen in order to circumscribe the
helical axis an increased number of times. A C3' endo pucker shortens the sugarphosphate backbone of DNA, bringing adjacent phosphates -1 A closer together than
they would be in the C2' endo form. Conversion of C3' endo moieties to the C2' endo
conformation allows the backbone to lengthen slightly in order to accommodate the
additional helix twist.
Positive and negative supercoiling produce a peak at 930 cm' and a trough at
1063 cm l'. Both are diagnostic of additional perturbations in DNA backbone structure.
The 930 cm -1 band has been assigned to a stretching vibration localized in the
deoxyribose ring and has been shown to decrease significantly in intensity upon binding
of the protein SRY to DNA (17). The intensity decrease was attributed to hydrophobic
contacts between SRY side chains and the deoxyribose rings of the DNA backbone. The
increased intensity at 930 cm-' observed in this study may be attributed to a change in
deoxyribose ring environments that occurs as a consequence of the significant bending
DNA undergoes when plectonemically supercoiled.
Comparison of Raman spectra of Supercoiled DNA with published values.
Several attempts have been made to ascertain the effects of supercoiling on DNA
structure by comparing the Raman spectrum of a supercoiled plasmid to either its relaxed
103
or linearized form (30-32). Such attempts have seen limited success, primarily due to
signal-to-noise ratios that are insufficient to reveal the subtle differences in structure
induced by supercoiling. Both instrumentation and spectral processing software have
improved considerably over time allowing for detection of more subtle changes in the
Raman spectrum of supercoiled DNA (33).
Most recently, the Raman spectrum of a negatively supercoiled pUC 19 plasmid
was compared to its linearized form (20). Interestingly, results from that study are not
consistent with the data presented here. The primary reason may be the difference in the
forms of DNA being compared. Comparing supercoiled to relaxed DNA will not
necessarily reveal the same spectral differences as comparing the supercoiled to the linear
form. Circularizing DNA necessarily imposes some restrictions on DNA geometry that
will dictate what conformations the molecule is free to sample. These restrictions may
manifest themselves in slight differences in the Raman spectrum of DNA, particularly in
the DNA backbone region. A recent study that compared Raman spectra of short (222
bp) predominately supercoiled circularized molecules with their linear pre-cursor found a
nearly null difference spectrum with modest bands at positions similar to those seen in
the supercoiled versus linear pUC19 study (34). Based on this finding, it seems likely
that the differences observed in the previous pUC 19 study better reflect the differences
between circular and linear DNA than the differences between supercoiled and relaxed
DNA. Comparison of relaxed pUC18 plasmid with the linear form may help clarify the
contributions of circularization versus supercoiling to the Raman spectrum.
104
CONCLUSIONS
In collaboration with colleagues at the University of Missouri at Kansas City, I
have generated Raman difference spectra to analyze the structural changes introduced to
the DNA by positive and negative supercoiling. When supercoiling of either direction is
introduced, the molecules remain in a predominately B-form conformation, with only
slight changes in structure occurring to compensate for the change in helix winding of the
DNA. These are primarily in the backbone of the molecule, with phosphodiester torsions
changing in both positively and negatively supercoiled DNA. Perhaps the most striking
observation was that of sugar pucker in positively supercoiled DNA converting to a more
homogenously C2'endo form to allow for the extra length in an overwound backbone.
Base stacking, was not significantly perturbed, but the environment of the exchangeable
protons of the bases was to some extent, suggesting a slight change in the accessibility of
the major groove as well as some destabilization of base pairing.
105
A
3
B
3.
C
D
400:
600
800
1000
1200
1400
1600
Figure 4.1. Raman spectra (300-1750 cm 1 752 nm excitation) of solutions of A) negatively
supercoiled and B) relaxed pUC18 plasmid DNA in 200 mM NaCl, pH 7.4 in H20 at 17 OC.
C) Difference spectrum between the two. D) Difference spectrum amplified x3.
106
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Figure 4.2. Raman spectra (300-1750 cm -1 532 nm excitation) of solutions of A) negatively
supercoiled and B) relaxed pUC18 plasmid DNA in 200 mM NaC1, pH 7.4 in H2 0 at 17 'C.
C) Difference spectrum between the two. D) Difference spectrum amplified x3.
107
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Figure 4.3. Raman spectra (300-1750 cm 1 752 nm excitation) of solutions of A) positively
supercoiled and B) relaxed pUC18 plasmid DNA in 200 mM NaCl, pH 7.4 in H2 0 at 17 'C.
C) Difference spectrum between the two. D) Difference spectrum amplified x3.
108
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532 nm excitation) of solutions of A) positively
Figure 4.4. Raman spectra (300-1750
supercoiled and B) relaxed pUC18 plasmid DNA in 200 mM NaC1, pH 7.4 in H20 at 17 oC.
C) Difference spectrum between the two. D) Difference spectrum amplified x3.
109
A
B
C
3-r
D
400
600
800
1000
12:00
1400
1600
Figure 4.5. Raman spectra (350-1800 cm 1' 752 nm excitation) of solutions of A) negatively
supercoiled and B) relaxed pUC18 plasmid DNA in 200 mM NaC1, pH 7.4 in D2 0 at 17 'C.
C) Difference spectrum between the two. D) Difference spectrum amplified x3.
110
A
B
ý3
2¸
C
D
400
600
800
1000
1200
1400
1600
Figure 4.6. Raman spectra (350-1750 cm 1 752 nm excitation) of solutions of A) positively
supercoiled and B) relaxed pUC18 plasmid DNA in 200 mM NaC1, pH 7.4 in D20 at 17 'C.
C) Difference spectrum between the two. D) Difference spectrum amplified x3.
111
LITERATURE CITATIONS
1.
Figueroa, N. and Bossi, L. (1988) Transcription induces gyration of the DNA
template in Escherichia coli. ProcNatl Acad Sci U S A, 85, 9416-9420.
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Brill, S.J. and Sternglanz, R. (1988) Transcription-dependent DNA supercoiling
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4.
Liu, L.F. and Wang, J.C. (1987) Supercoiling of the DNA template during
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Funnell, B.E., Baker, T.A. and Kornberg, A. (1986) Complete enzymatic
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Droge, P. (1993) Transcription-driven site-specific DNA recombination in vitro.
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Thomas, G.J. (1999) Raman spectroscopy of protein and nucleic acid assemblies.
Annu Rev Biophys Biomol Struct, 28, 1-27.
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Benevides, J.M. and Thomas, G.J., Jr. (1983) Characterization of DNA structures
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Benevides, J.M., Weiss, M.A. and Thomas, G.J., Jr. (1991) Design of the helixturn-helix motif: nonlocal effects of quaternary structure in DNA recognition
investigated by laser Raman spectroscopy. Biochemistry, 30, 4381-4388.
13.
Benevides, J.M., Weiss, M.A. and Thomas, G.J., Jr. (1991) DNA recognition by
the helix-turn-helix motif: investigation by laser Raman spectroscopy of the phage
112
lambda repressor and its interaction with operator sites OL 1 and OR3.
Biochemistry, 30, 5955-5963.
14.
Benevides, J.M., Wang, A.H., van der Marel, G.A., van Boom, J.H. and Thomas,
G.J., Jr. (1989) Effect of the G.T mismatch on backbone and sugar conformations
of Z-DNA and B-DNA: analysis by Raman spectroscopy of crystal and solution
structures of d(CGCGTG) and d(CGCGCG). Biochemistry, 28, 304-310.
15.
Benevides, J.M. and Thomas, G.J., Jr. (2005) Local conformational changes
induced in B-DNA by ethidium intercalation. Biochemistry, 44, 2993-2999.
16.
Benevides, J.M., Li, T., Lu, X.J., Srinivasan, A.R., Olson, W.K., Weiss, M.A. and
Thomas, G.J., Jr. (2000) Protein-directed DNA structure II. Raman spectroscopy
of a leucine zipper bZIP complex. Biochemistry, 39, 548-556.
17.
Benevides, J.M., Chan, G., Lu, X.J., Olson, W.K., Weiss, M.A. and Thomas, G.J.,
Jr. (2000) Protein-directed DNA structure. I. Raman spectroscopy of a highmobility-group box with application to human sex reversal. Biochemistry, 39,
537-547.
18.
Benevides, J.M., Kang, C. and Thomas, G.J., Jr. (1996) Raman signature of the
four-stranded intercalated cytosine motif in crystal and solution structures of
DNA deoxycytidylates d(CCCT) and d(C8). Biochemistry, 35, 5747-5755.
19.
Benevides, J.M., Wang, A.H., van der Marel, G.A., van Boom, J.H., Rich, A. and
Thomas, G.J., Jr. (1984) The Raman spectra of left-handed DNA oligomers
incorporating adenine-thymine base pairs+. Nucleic Acids Res, 12, 5913-5925.
20.
Serban, D., Benevides, J.M. and Thomas, G.J., Jr. (2002) DNA secondary
structure and Raman markers of supercoiling in Escherichia coli plasmid pUC 19.
Biochemistry, 41, 847-853.
21.
Prescott, B., Steinmetz, W. and Thomas, G.J., Jr. (1984) Characterization of DNA
structures by laser Raman spectroscopy. Biopolymers, 23, 235-256.
22.
Erfurth, S.C. and Peticolas, W.L. (1975) Melting and premelting phenomenon in
DNA by laser Raman scattering. Biopolymers, 14, 247-264.
23.
Movileanu, L., Benevides, J.M. and Thomas, G.J., Jr. (2002) Determination of
base and backbone contributions to the thermodynamics of premelting and
melting transitions in B DNA. Nucleic Acids Res, 30, 3767-3777.
24.
Toyama, A., Takeuchi, H. and Harada, I. (1991) Ultraviolet resonance Raman
spectra of adenine, uracil and thymine derivatives in several solvents. Correlation
between band frequencies and hydrogen-bonding states of the nucleic acid bases.
Journalof Molecular Structure, 242, 87-98.
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25.
Movileanu, L., Benevides, J.M. and Thomas, G.J. (1999) Temperature
dependence of the Raman spectrum of DNA. Part I - Raman signatures of
premelting and melting transitions of Poly(dA-dT).Poly(dA-dT). Journalof
Raman Spectroscopy, 30, 637-649.
26.
Deng, H., Bloomfield, V.A., Benevides, J.M. and Thomas, G.J., Jr. (1999)
Dependence of the Raman signature of genomic B-DNA on nucleotide base
sequence. Biopolymers, 50, 656-666.
27.
Benevides, J.M., Kukolj, G., Autexier, C., Aubrey, K.L., DuBow, M.S. and
Thomas, G.J. (1994) Secondary Structure and Interaction of Phage D108 Ner
Repressor with a 61-Base-Pair Operator: Evidence for Altered Protein and DNA
Structures in the Complex. Biochemistry, 33, 10701-10710.
28.
Thomas, G.J., Jr. and Benevides, J.M. (1985) An A-helix structure for poly(dAdT) X poly(dA-dT). Biopolymers, 24, 1101-1105.
29.
Westhof, E. (1987) Re-refinement of the B-dodecamer d(CGCGAATTCGCG)
with a comparative analysis of the solvent in it and in the Z-hexamer
d(5BrCG5BrCG5BrCG). JBiomol Struct Dyn, 5, 581-600.
30.
Christens-Barry, W.A., Martin, J.C. and Lebowitz, J. (1989) Raman spectroscopy
of supercoiled and nicked ColEl plasmid. Biopolymers, 28, 1515-1526.
31.
Brahms, S., Nakasu, S., Kikuchi, A. and Brahms, J.G. (1989) Structural changes
in positively and negatively supercoiled DNA. Eur JBiochem, 184, 297-303.
32.
Vasmel, H. (1985) Influence of supercoiling on DNA structure: laser Raman
spectroscopy of the plasmid pBR322. Biopolymers, 24, 1001-1008.
33.
Benevides, J.M., Overman, S.A. and Thomas, G.J. (2005) Raman, polarized
Raman and ultraviolet resonance Raman spectroscopy of nucleic acids and their
complexes. JRaman Spectroscopy, 36, 279-299.
34.
Benevides, J.M., Serban, D. and Thomas, G.J., Jr. (2006) Structural perturbations
induced in linear and circular DNA by the architectural protein HU from Bacillus
stearothermophilus. Biochemistry, 45, 5359-5366.
114
Chapter 5 - Design and Optimization of Fluorescent DNA Substrates
for Micromanipulator Studies
INTRODUCTION
In previous chapters, I have demonstrated that physiological levels of positive
supercoiling have subtle effects on the structure of DNA. Both positive and negative
supercoiling are accommodated by small changes in sugar-phosphate backbone and a low
level disruption of base pairing. While crude chemical and enzymatic probes showed no
evidence of base unpairing in positively supercoiled DNA, Raman spectral data indicate
that disruption of pairing does indeed occur in a small fraction of the bases in positively
supercoiled molecules, accompanied by slight changes in phosphodiester torsion and
sugar pucker. These studies, however, were limited to plasmid substrates with a fixed
level of superhelical density. Just as DNA itself is a dynamic molecule, supercoiling is a
dynamic parameter, and subject to a number of determining factors. Topoisomerase
activity, level of transcription, and relative orientation of neighboring genes in the same
topological domain will all affect the degree of supercoiling present (1,2). Additionally,
supercoiling is not always evenly distributed throughout a given domain; supercoiling
generated by an actively processing enzyme will exist in a gradient, with the highest
degree of supercoiling immediately adjacent to the enzyme (3). Because the degree of
supercoiling can vary so greatly, it would be useful to examine its consequences at a wide
range of superhelical densities.
115
Since plasmids are usually examined at a fixed superhelical state, they only
provide information at one point along the continuum. While it is possible to isolate
individual topoisomers for experimentation, obtaining the requisite amounts for detailed
analysis is difficult, and technical limitations prevent the overall level of positive
supercoiling from being above a certain superhelical density. Therefore, a potentially
more informative line of experimentation would involve a substrate in which the
superhelical density could be easily manipulated and precisely controlled over a wide
range of superhelical densities.
One approach to this level of precise control involves the use of a magnetic
micromanipulator. This entails tethering one end of a surface-anchored DNA molecule
to a magnetic bead that is manipulated in a controlled magnetic field, which allows for
incremental rotation of the DNA helix (4-6). Such studies thus far, however, have only
looked at global changes in the molecular structure after phase transition, rather than
elucidating the fine structural alterations that could occur with lower levels of introduced
torsion (4-6). The nature of the data obtained in these earlier studies precludes such fine
detail, as the only parameters measured are the force applied to the bead, the length of the
DNA, and the superhelical density (4-6).
Considerably more information could be obtained by including a sensitive
reporter of DNA structure. One such reporter is 2-aminopurine (2AP), which is a
fluorescent base analog widely used in oligonucleotide-based studies of DNA structure
and DNA-protein interactions (7-11). 2AP forms a stable base pair with thymine, in a
Watson-Crick geometry (12) that is only slightly less stable than its A:T counterpart (Fig
5.1) (13). It has an excitation maximum at 303 nm, allowing it to be selectively excited
116
at a wavelength that will reduce autofluorescence from canonical bases in the helix (14).
Perhaps most importantly, fluorescence properties of 2AP are extremely sensitive to the
environment of the base: Both the fluorescence yield and lifetime are significantly
reduced when the base analog is stacked in a normal B-form helix, and the effects on the
fluorescence properties for various perturbations in the environment of the base have
been studied in some detail (7,8,14,15). Thus, 2AP has been used as a sensitive probe to
investigate changes in the base environment resulting any number of factors, ranging
from the effects of abasic sites on neighboring bases (7,8), to the action of "baseflipping" enzymes that act by everting a base from the DNA helix into an active site
pocket of the protein (9,16).
If positive supercoiling leads to solvent exposure of DNA bases, then 2AP should
help reveal the nature and extent of this exposure. Combining 2AP-containing DNA with
a magnetic micromanipulator would allow for structural changes to be determined as a
function of twist, providing a detailed picture of the base environment as precisely
controlled amounts of twist are introduced. This would significantly broaden our
understanding of overwinding and its effect on the solvent environment of DNA bases.
In this chapter, I describe preliminary efforts to prepare 2-aminopurine-containing
micromanipulator substrates. Current technical limitations have rendered these studies
unfeasible, but the information obtained in these preliminary efforts should prove
valuable for future efforts at studying changes to the base environment using a
micromanipullation approach.
117
MATERIALS AND METHODS
Design of a 2-aminopurine-containingDNA substratefor use in a magnetic
micromanipulator. A detailed description of the design for the micromanipulator
substrate is given in Figures 5.2 and 5.3. Briefly, a 2.1 kb segment of 2-aminopurinecontaining DNA is generated using PCR, and digested with restriction enzymes to
produce unique overhangs on each end. These fragments are dimerized, and ligated to a
large X-DNA restriction fragment to produce a circular molecule. A further restriction
digest removes a small fragment of the circle opposite the 2AP-labeled portion, resulting
in a long linear molecule with a region of incorporated 2AP in the middle. Short linker
fragments, generated by restriction digestion of plasmid pUC 18, followed by labeling
with either biotin or digoxygenin, are then ligated on to the ends of the long molecule,
and the resultant substrate is ready for use in the micromanipulator.
Introduction of2-aminopurine into DNA substrates using modified PCR. A 2135
bp PCR product was amplified from plasmid pBR322 using the primers 5'CCTGCTCGCTTCGCTACTTG-3' and 5'-GGATAACCGTATTACCGCCTTTG-3'
with Taq polymerase (New England Biolabs, Ipswich, MA). Reactions were carried out
in lx ThermoPol Buffer (20 mM Tris-HC1, 10 mM (NH 4 )2 SO4 , 10 mM KC1, 2 mM
MgSO 4 , 0.1% Triton X- 100, pH 8.8). 2-Aminopurine-2' deoxyribose-5'-triphosphate
(dAPTP) (TriLink Biotechnologies, San Diego, CA) was added to the reaction mixture to
a concentration of 150 tiM; the concentration of all other dNTPs was 200 rtM. After an
initial denaturation step at 95 'C for 5 min, 30 amplification cycles were performed,
118
consisting of the following segments: 95 TC for 30 sec, 61 TC for 60 s and 72 TC for 105
s. Following the amplification cycles, a final elongation step at 72 'C for 7 min was
performed. 2-AP incorporation was assessed by fluorescence emission yield (ex: 303
nm, em: 370 nm) after purification and depurination.
Digestion andpurification of componentsfor the micromanipulatorsubstrate.
Restriction enzyme digests to obtain the requisite DNA fragments are summarized in
Table 5.1. Following digestion, the desired fragments were isolated by resolution on a
Gen-Pak FAX anion exchange column (Waters, Milford, MA), using an Agilent 1100
HPLC (Santa Clara, CA). Fragments were eluted using a gradient of 0 to 1 M NaCl in
TE buffer (10 mM Tris, pH 8.0, 1 mM EDTA) at a flow rate of 0.75 mL/min for 30 min.
Absorbance at 254 nm was used to identify peaks for collection, which were then
precipitated by addition of an equal volume of ice-cold isopropanol, followed by
centrifugation at 16,000 x g for 30 min. The resulting pellets were air dried and
resuspended in TE prior to further manipulation.
Labeling of linkerfragments with biotin or digoxygenin. Following HPLC
purification, linker segments of DNA were labeled with either biotin or digoxygenin
using Chem-Link labeling kits (Roche Applied Sciences, Indianapolis, IN) according to
the instructions provided, with the exception of a reduced reaction temperature of 65 'C.
Degree of labeling was assessed colorimetrically using streptavidin- or anti-DIG-linked
alkaline phosphatase in conjunction with NBT/BCIP (Roche Applied Sciences).
Fluorescence lifetime studies of 2-aminopurine substrates in solution. For
fluorescence lifetime measurements of both free 2APTP and 2AP-containing DNA,
samples were excited with femtosecond pulses from a Ti:sapphire laser (Mira, Coherent,
119
Santa Clara, CA) tuned to 909 nm and frequency tripled to 303 nm with an Inrad model
5-050 frequency harmonic generator (Northvale, NJ). Emission was detected at 370 nm
using a custom-made microscope based on an Axiovert 110 inverted microscope (Zeiss,
G6ttingen, Germany) equipped with a Becker-Hickl 730 time-correlated single photon
counting module (Berlin, Germany). Steady state total internal reflection (TIR)
fluorescence was measured using the same instrumentation; the beam was reflected in a
quartz prism to produce the evanescence required for excitation, and the emission was
measured using the inverted microscope described above equipped with a photomultiplier
tube (R5600-P, Hamamatsu, Bridgewater, NJ).
RESULTS
2-Aminopurine can be efficiently incorporatedinto DNA using PCR. 2Aminopurine incorporation has frequently been used to study polymerase fidelity (1719), however, a PCR-based method to generate densely-labeled 2-aminopurinecontaining substrates has previously been lacking. Given the stable base pairing of 2AP
with thymine (12), and the high level of incorporation by some polymerases (20,21), it is
reasonable to expect that such a method could be established. In fact, adding 2AP
deoxynucleoside triphosphates to the PCR reaction mixture leads to significant levels of
incorporation, under appropriate conditions.
In recent years, considerable effort has been directed toward engineering
thermostable polymerases with improved fidelity. In contrast, our needs demand a
polymerase with relatively low fidelity and also limited 3' -5' exonuclease activity (21).
120
By using a polymerase with a higher error rate and limited proofreading capabilities, the
amount of incorporated 2AP can be maximized.
While increasing the concentration of 2AP-containing dNTP (dAPTP) increases
the level of incorporation, the trade-off is that the overall product yield is diminished.
Therefore, a 150 [tM concentration of dAPTP was selected for preparation of 2APcontaining substrates, as it provides a reasonable level of incorporation (-1 in every 50
bp), while still resulting in acceptable product yield. Having enough 2AP bases
incorporated is crucial for adequate detection, but excessive 2AP labeling may partially
destabilize the helix, and would thus be undesirable.
Buffer conditions are critical for efficient incorporation of dAPTP by the
polymerase. While 2AP was well-incorporated when using New England Biolabs
ThermoPol buffer (20 mM Tris-HC1, 10 mM (NH4 )2 SO 4 , 10 mM KC1, 2 mM MgSO 4 ,
0.1% Triton X- 100; pH 8.8), the same company's Standard Taq Reaction buffer (10 mM
Tris-C1, 50 mM KC1, 1.5 mM MgCl 2 ; pH 8.3) resulted in no detectable 2AP
incorporation. The reasons for this were not fully investigated, although an increase in
Mg 2+ concentration and a increase in pH have both been demonstrated to reduce the
fidelity of Taq polymerase (22).
Functionalizationof linker DNA with Chem-Link labeling kits. To retain
introduced torsion, both strands of the DNA substrate must be securely anchored, thus
multiple biotins distributed on each strand of the linker are desirable. Two methods of
biotinylation of the linker DNA were tested: EZ-Link photoactivatable biotin (Pierce
Biotechnology, Rockford, IL), and Chem-Link biotin (Roche Applied Science). The
latter was determined to be considerably more effective, with a greater than ten-fold
121
increase in labeling, therefore Chem-Link was also used to functionalize linker DNA
with digoxygenin.
PurificationofDNA components using HPLC The Gen-Pak Fax column was
effective in purifying desired components ranging in size from -800 to -18000 (23).
Restriction digests were optimized so as to maximize the size differential between the
desired component and the remaining fragments, so that the separations were clean and
effective in all cases. Theoretically, fragments that are closer in size than those purified
in this study could be resolved by slightly adjusting the NaCl gradient, but this was not
necessary in any of these separations. Elution of fragments with NaCl in TE buffer
facilitates further manipulation, as eluted fragments can be easily purified with an alcohol
precipitation step.
Fluorescence lifetime offree and incorporated2-aminopurine. As an initial test
of both the instrumentation and the DNA substrate, the fluorescence lifetimes of free
dAPTP and 2AP incorporated into purified PCR product were measured in solution (Fig.
5.4). The fluorescence lifetime of free dAPTP was 10.9 ns, consistent with previously
published values (7,24). The lifetime for the incorporated 2AP had both a fast and a slow
decay component: 55% 0.13 ns, and 45% 3.28 ns. Other values for quenched 2AP
lifetimes were in line with these values, although most studies use more lifetime
components to fit the data (7,24-26). Any variance from previously published values is
likely because those values reflect 2AP incorporation at a single location within an
oligonucleotide, while the PCR product has 2AP incorporation into multiple contexts
opposite a thymine base; sequence context is known to affect fluorescence properties of
2AP in DNA (15). Despite these small differences, this result clearly shows that 2-
122
aminopurine is incorporated sufficiently into DNA using PCR, and that fluorescence
signal quenched to a degree that will allow for easy discernment of bases that are
normally stacked and paired relative to extrahelical bases.
Preliminarytests of2-aminopurine in the TIR instrumentation. In order to
measure sensitivity of the micromanipulator's optical instrumentation prior to assembly
of the substrate within the system, dilutions of dAPTP were tested for signal levels.
Despite an early calculation suggesting that a 20 nM effective concentration would be
sufficient for all proposed experiments, testing showed that a 1 [tM solution would be
necessary for a 2:1 signal-to-noise ratio of fluorescence emission.
In order to calculate the effect of this difference on the feasibility of our system,
we first had to consider the volume of excitation. The laser beam exciting the TIR has a
diameter of 3x10-3 m, for a total area of 7.1x10 -6 m. The evanescent waves that excite the
substrate in TIR are strongest at the interface between the reflective surface and the
solvent medium, and decrease exponentially as the distance from the interface increases.
For the quartz prism used, we will assume an effective excitation depth of 120 nm,
keeping in mind that the level of excitation will not be homogenous throughout. This
gives us a solution volume of 8.5x10-10 L. Since a 1 tM solution is necessary for
adequate signal, a total of 8.5x10-16 mol of unquenched 2AP is necessary to have a
detectable fluorescence signal.
We can now compare this value to the number of moles of 2AP that we could
reasonably expect within our proposed DNA substrates. The magnetic beads used in
these experiments are 1 x 10-6m in diameter, so the maximum expected labeling of the
surface with DNA strands would be approximately one strand every 2 x 10-6 m 2. In the
123
excitation area of the solvent medium, we could then expect to have roughly 3.5 x 106
DNA molecules.
Assuming optimal 2AP labeling, we could expect approximately one 2AP base
for every 50 bp of DNA. With a B-form DNA rise of 0.34 nm per bp, roughly the first
350 bp of every strand would be excited. This gives an approximate total of 7 2AP
residues per molecule, for a total of 4.2x1 017 molecules of 2AP within the excitation
volume.
In the case of unquenched fluorophore, this is roughly 20-fold fewer 2AP
molecules than would be necessary for adequate signal, but our experiments require that
we measure signal from paired and stacked fluorophores as well as from unquenched
molecules. 2AP is quenched significantly when it is incorporated into B-form DNA,
between 25- to 125-fold, depending on the context (10). If the 2AP is only quenched 25fold within the context of the helix, the described system would fail to detect the full
range of 2AP, from completely stacked to completely extrahelical, by approximately 500fold.
Even assuming ideal conditions, the fluorescence yield is significantly less than
that required for viable experiments. Incorporation of 2AP and the density of DNA
strands on the surface are near to their maximal values, therefore other changes must be
made in order to realize the proposed studies.
124
DISCUSSION
In this study, I have made initial efforts to develop a system that combines an
incorporated fluorescent base analog with a magnetic micromanipulator in order to study
changes in the DNA base environment that result from supercoiling. These experiments
are not yet technically feasible; however, important considerations were revealed which
will aid any future efforts. Most notable among these are the development of a method to
incorporate fluorescent bases into DNA substrates using PCR, and the optimization of
substrate design for use in a fluorescence micromanipulator system.
The fluorescent base analog 2-aminopurine can be readily incorporated into PCR
substrates using appropriate solvent conditions and a suitably low-fidelity polymerase.
However, despite the utility of 2AP in oligonucleotide-based studies of DNA structure, it
may not be ideal for use in a TIR excitation system. 2AP excites in the low-UV range,
leading to considerable autofluorescence from any impurities in the solvent and in the
quartz prism, as well as a low fluorescence efficiency and a high rate of photobleaching.
Choosing a fluorophore with a red-shifted excitation maximum could significantly reduce
these problems.
One such fluorophore is 6-methylisoxanthopterin (6MI), a bicyclic pteridine
analog that pairs with cytosine and has an excitation maximum at 340 nm (27). The
quantum yield is similar to 2AP: 0.70 (27) compared to 0.68 for 2AP (14). Both the
quantum yield and fluorescence lifetime of 6MI are significantly reduced upon
incorporation into a DNA helix, although this effect is less pronounced than it is for 2AP
(10,27). Base: pairing is stable with 6MI; only slight melting point depression relative to
125
the unsubstituted form was observed when 6MI was incorporated into an oligonucleotide
opposite a cytosine (27). Additionally, a recent study has shown 6MI to be amenable to
two-photon excitation, while three-photon excitation of 2AP was significantly less viable
(28).
The fluorescence properties of 6MI, particularly with an excitation wavelength
that is red-shifted relative to that of 2AP, make it an intriguing candidate for use in
micromanipulator studies. However, two major obstacles exist to testing 6MI for these
purposes: incorporation and availability. While both 2AP and 6MI form Watson-Crick
base pairs, 2AP is a purine rather than a pteridine, and as such may be more readily
incorporated by a polymerase. It is possible that a PCR-based method could be
developed to incorporate 6MI into DNA, however this may prove difficult, and would
require further investigation. Additionally, 6MI has thus far seen limited use, and is not
now commercially available in base, nucleoside, or nucleotide form, so obtaining
sufficient quantities to construct a substrate is likely to be either expensive or laborintensive.
In addition to the possible use of a fluorophore, this study has also provided
insights into effective substrate design for use in a fluorescence-based micromanipulation
system. The most apparent change that arises in substrate design is the location of the
fluorophore-containing DNA segment. The original substrate had the 2AP-containing
portion in the middle, flanked by two pieces of unlabeled extender DNA, which are then
attached to the linkers (Fig. 5.5a). Because TIR excitation is effective only within a
limited distance from the surface of the reflective prism, the fluorophore-containing
region should be as close to that surface as possible. This means that the 2AP-containing
126
fragment should be connected directly to surface linker segment, with a single long piece
of unlabeled extender DNA connected to the bead-linker region (Fig. 5.5b). In addition
to providing better excitation of the fluorophore, such a substrate would be considerably
simpler to construct, as it would require fewer ligation and purification steps.
Another important consideration arising from our initial experiments is the
distance between the fluorophore-containing portion of the DNA molecule and the
excitation source, which is determined by the length of the linker region. If TIR is to be
used as an excitation source, the fluorophores in the DNA molecule must be located as
close to the solid support as possible for maximal fluorescence yield. In the original
substrate, there were two significant problems that must be rectified.
The first involves is the length of the linker, since the longer it is, the further from
the excitation source any attached fluorophore-containing DNA will be. With the current
linker length of -800 bp, the linker DNA would be approximately 275 nm in length.
While the linker will be flattened against the surface to some extent due to contacts
between the functional groups on the linker and their conjugates on the surface, some of
the linker DNA is expected to be free from these linkages and will therefore contribute to
the overall distance between the excitation source and the DNA containing the
fluorophores. Minimizing the length of this linker DNA is therefore desirable.
The second problem with the initial design was the use of digoxygenin (DIG)
bound to anti-DIG antibodies as a method for attaching the linker to the quartz prism. In
this scheme, the DNA was functionalized with DIG, and the prism surface was coated
with anti-DIG antibodies. This linkage will both interfere with excitation of the
fluorophore, and add to the noise in the system. In terms of interference of excitation,
127
IgG antibodies are on the order of 15 nm in total length (29), meaning that the linker
would not be immediately proximal to the surface, but instead would be offset by the
equivalent of -44 bp in length. Since the evanescence in the current system is only
effective up to -120 nm, and is strongest at the prism surface, a significant fraction of the
excitation energy will not reach the DNA. Additionally, with antibodies coating the TIR
interface, the strongest available evanescence will excite the weakly fluorescent amino
acids in the protein, leading to a high background signal.
To eliminate these problems, the length of the linker used should be shortened,
and a linkage scheme that does not employ antibodies should be used. The linkage used
to connect the DNA substrate to the magnetic bead is not subject to such limiting
requirements, so it may be possible to swap linkages and use the DIG-anti-DIG for
connecting to the bead while using smaller molecules, biotin and streptavidin, to connect
to the solid support. Alternatively, covalent linkages could be designed to directly anchor
both strands to the surface. In either case, a short oligonucleotide with the appropriate
functionalization on each strand should suffice. The main requirements for any linker
would remain the same: it needs to bind both strands of the DNA to the support with
sufficient strength to allow retention of applied torsion, and it must not significantly
perturb the structure of the DNA.
The realization of a magnetic micromanipulator system in which alterations to the
base environment can be measured fluorometrically as a function of twist will likely
require significantly more optimization, both of the instrumentation and the DNA
substrates. Still, this would further our understanding of supercoiling-induced changes to
128
DNA structure tremendously. Information gained from this preliminary study will serve
as a guide for any future studies.
129
H
-n
,,
H3
4N
N
N
-/
n'
'
H13
N
/P/N
---- H///ý-N
N
/ N-H
----0O
2N
H
N
----
,+N-H
N
N
N•
_•H
N--H ----
0
H
Figure 5.1. Hydrogen bonding schemes of A) the canonical Watson-Crick adenine-thymine
base pair B) the fluorescent base analog 2-aminopurine (2AP) paired to thymine, and C) a
proposed protonated form of 2AP that could pair with cytosine.
130
*If
~*I
MtI,`IMLý
Figure 5.2. Preparation of components of micromanipulator substrate. A) A 2AP-containing
PCR product is amplified from pBR322, and digested with BamHI and ApaLI to yield a
fragment with different four base 5'-overhangs on each end. B) Linker fragments of -800 bp
are prepared by double digestion of pUC18 with either XmnI/XbaI or XmnI/BamHI. Linkers
were labeled with either biotin or digoxygenin using Roche Chem-Link labeling reagents. C)
An 18,909 bp fragment is isolated from a BclI digestion of N6-methyladenine-free X-DNA,
and treated with BamHI methyltransferase to protect the two BamHI recognition sites from
cleavage in subsequent steps.
131
1
-
~+-
I
I
M
.9,,
.4
qyly
Figure 5.3. Assembly scheme for micromanipulator substrate. A) 2-aminopurine-containing
fragments are ligated in the presence of BamHI, generating dimers with identical four base
5'overhangs on each end. B) These dimers are ligated to the X-DNA fragment in the presence
of BamHI and BclI to prevent self-ligation, in dilute conditions to promote circle formation.
C) These circles are double digested with BglII and Xbal to form a linear molecule with
unique four base 5'overhangs. D) Labeled linker DNA is ligated to the ends of the linear
molecule, completing the substrate.
132
0.75
U
C
U
U)
o
:3
9
0.5
V
N
E
z
0
o
ZC
0.25
0
1
2
3
4
5
6
7
Time (ns)
Figure 5.4. Fluorescence lifetime of free (top curve) and incorporated (bottom curve) 2aminopurine. Lifetime was significantly quenched by incorporation into the DNA helix.
Fluorescence lifetimes were determined to be: free - 10.9 ns, incorporated - 55% 0.13 ns, 45%
3.28 ns.
133
B
•1111
•111
Figure 5.5. Comparison of original design of micromanipulator substrate with optimized
design. A) Substrate as originally designed. Fluorophore is located in the middle of the
substrate, too distant from the quartz surface to be excited. Antibodies link a long linker to
the surface, putting the fluorophore containing region at a further distance from the excitation.
B) Substrate design optimized for TIR excitation. Fluorophore-containing region is as close
to the quartz surface as possible, linked to the surface by a short, non-antibody-based linker.
134
LITERATURE CITATIONS
1.
Wu, H.Y., Shyy, S.H., Wang, J.C. and Liu, L.F. (1988) Transcription generates
positively and negatively supercoiled domains in the template. Cell, 53, 433-440.
2.
Tsao, Y.P., Wu, H.Y. and Liu, L.F. (1989) Transcription-driven supercoiling of
DNA: direct biochemical evidence from in vitro studies. Cell, 56, 111-118.
3.
Wang, Z. and Droge, P. (1997) Long-range effects in a supercoiled DNA domain
generated by transcription in vitro. J Mol Biol, 271, 499-510.
4.
Allemand, J.F., Bensimon, D., Lavery, R. and Croquette, V. (1998) Stretched and
overwound DNA forms a Pauling-like structure with exposed bases. Proc Natl
Acad Sci USA, 95, 14152-14157.
5.
Strick, T.R., Allemand, J.F., Bensimon, D., Bensimon, A. and Croquette, V.
(1996) The elasticity of a single supercoiled DNA molecule. Science, 271, 18351837.
6.
Strick, T.R., Allemand, J.F., Bensimon, D. and Croquette, V. (1998) Behavior of
supercoiled DNA. Biophys J, 74, 2016-2028.
7.
Rachofsky, E.L., Seibert, E., Stivers, J.T., Osman, R. and Ross, J.B. (2001)
Conformation and dynamics of abasic sites in DNA investigated by time-resolved
fluorescence of 2-aminopurine. Biochemistry, 40, 957-967.
8.
Stivers, J.T. (1998) 2-Aminopurine fluorescence studies of base stacking
interactions at abasic sites in DNA: metal-ion and base sequence effects. Nucleic
Acids Res, 26, 3837-3844.
9.
McCullough, A.K., Dodson, M.L., Scharer, O.D. and Lloyd, R.S. (1997) The role
of base flipping in damage recognition and catalysis by T4 endonuclease V. J Biol
Chem, 272, 27210-27217.
10.
Bloom, L.B., Otto, M.R., Eritja, R., Reha-Krantz, L.J., Goodman, M.F. and
Beechem, J.M. (1994) Pre-steady-state kinetic analysis of sequence-dependent
nucleotide excision by the 3'-exonuclease activity of bacteriophage T4 DNA
polymerase. Biochemistry, 33, 7576-7586.
11.
Raney, K.D., Sowers, L.C., Millar, D.P. and Benkovic, S.J. (1994) A
fluorescence-based assay for monitoring helicase activity. Proc Natl Acad Sci US
A,91, 6644-6648.
12.
Sowers, L.C., Fazakerley, G.V., Eritja, R., Kaplan, B.E. and Goodman, M.F.
(1986) Base pairing and mutagenesis: observation of a protonated base pair
between 2-aminopurine and cytosine in an oligonucleotide by proton NMR. Proc
Natl Acad Sci USA, 83, 5434-5438.
135
13.
Eritja, R., Kaplan, B.E., Mhaskar, D., Sowers, L.C., Petruska, J. and Goodman,
M.F. (1986) Synthesis and properties of defined DNA oligomers containing base
mispairs involving 2-aminopurine. Nucleic Acids Res, 14, 5869-5884.
14.
Ward, D.C., Reich, E. and Stryer, L. (1969) Fluorescence studies of nucleotides
and polynucleotides. I. Formycin, 2-aminopurine riboside, 2,6-diaminopurine
riboside, and their derivatives. JBiol Chem, 244, 1228-1237.
15.
Jean, J.M. and Hall, K.B. (2002) 2-Aminopurine electronic structure and
fluorescence properties in DNA. Biochemistry, 41, 13152-13161.
16.
Allan, B.W., Beechem, J.M., Lindstrom, W.M. and Reich, N.O. (1998) Direct real
time observation of base flipping by the EcoRI DNA methyltransferase. JBiol
Chem, 273, 2368-2373.
17.
Goodman, M.F., Hopkins, R. and Gore, W.C. (1977) 2-Aminopurine-induced
mutagenesis in T4 bacteriophage: a model relating mutation frequency to 2aminopurine incorporation in DNA. Proc Natl Acad Sci US A, 74, 4806-4810.
18.
Clayton, L.K., Goodman, M.F., Branscomb, E.W. and Galas, D.J. (1979) Error
induction and correction by mutant and wild type T4 DNA polymerases. Kinetic
error discrimination mechanisms. JBiol Chem, 254, 1902-1912.
19.
Watanabe, S.M. and Goodman, M.F. (1982) Kinetic measurement of 2aminopurine X cytosine and 2-aminopurine X thymine base pairs as a test of
DNA polymerase fidelity mechanisms. Proc Natl Acad Sci USA, 79, 6429-6433.
20.
Grossberger, D. and Clough, W. (1981) Incorporation into DNA of the base
analog 2-aminopurine by the Epstein-Barr virus-induced DNA polymerase in vivo
and in vitro. ProcNatl Acad Sci US A, 78, 7271-7275.
21.
Pless, R.C., Levitt, L.M. and Bessman, M.J. (1981) Nonrandom substitution of 2aminopurine for adenine during deoxyribonucleic acid synthesis in vitro.
Biochemistry, 20, 6235-6244.
22.
Eckert, K.A. and Kunkel, T.A. (1990) High fidelity DNA synthesis by the
Thermus aquaticus DNA polymerase. Nucleic Acids Res, 18, 3739-3744.
23.
Strege, M.A. and Lagu, A.L. (1991) Anion-exchange chromatography of DNA
restriction fragments. J Chromatogr,555, 109-124.
24.
Guest, C.R., Hochstrasser, R.A., Sowers, L.C. and Millar, D.P. (1991) Dynamics
of mismatched base pairs in DNA. Biochemistry, 30, 3271-3279.
25.
Rachofsky, E.L., Osman, R. and Ross, J.B. (2001) Probing structure and
dynamics of DNA with 2-aminopurine: effects of local environment on
fluorescence. Biochemistry, 40, 946-956.
136
26.
Nordlund, T.M., Andersson, S., Nilsson, L., Rigler, R., Graslund, A. and
McLaughlin, L.W. (1989) Structure and dynamics of a fluorescent DNA oligomer
containing the EcoRI recognition sequence: fluorescence, molecular dynamics,
and NMR studies. Biochemistry, 28, 9095-9103.
27.
Hawkins, M.E., Pfleiderer, W., Balis, F.M., Porter, D. and Knutson, J.R. (1997)
Fluorescence properties of pteridine nucleoside analogs as monomers and
incorporated into oligonucleotides. Anal Biochem, 244, 86-95.
28.
Katilius, E. and Woodbury, N.W. (2006) Multiphoton excitation of fluorescent
DNA base analogs. JBiomed Opt, 11, 044004.
29.
Werner, T.C., Bunting, J.R. and Cathou, R.E. (1972) The shape of
immunoglobulin G molecules in solution. Proc Natl Acad Sci US A, 69, 795-799.
137
Chapter 6 - Effects of Supercoiling on Cleavage by Type II Restriction
Enzymes
INTRODUCTION
In previous chapters, I have addressed structural changes that occur with the
introduction of positive supercoiling into DNA. Both positively and negatively
supercoiled DNA accommodate introduced torsion with small conformational changes in
base pairing and the sugar-phosphate backbone. Details from Raman difference
spectroscopy suggest that with positively supercoiled DNA these changes include altered
phosphodiester torsion, sugar pucker, and base pairing status. It is crucial to elucidate
these structural transitions in order to understand the interaction of DNA with other
cellular constituents, including proteins and small molecules. Since unconstrained
supercoiling generated by processive enzymes is ubiquitous in the DNA of both
prokaryotes and eukaryotes (1-3), proteins that interact with DNA are likely to encounter
supercoiling in the course of normal cellular activity. Even with subtle changes in DNA
structure, supercoiling has been shown to alter the way in which proteins are able to
interact with DNA, through mechanisms stemming from both the twist and writhe
components (4,5).
While: many proteins have been identified that have differential interactions with
negatively supercoiled DNA (6-10), positively supercoiled DNA has not been as wellexamined, though some examples of altered binding or reactivity are known (8,9,11).
139
Enzymes that may be active on both positively and negatively supercoiled substrates are
likely to interact with each substrate in subtly different ways.
One example is human topoisomerase I, a Type IB topoisomerase that nicks one
strand of DNA and allows elastic free energy to drive controlled rotation around the
intact strand, thus relieving superhelical tension (12). The crystal structure of the
substrate-bound protein shows the enzyme as a hinged structure with two "caps", which
wrap around the substrate DNA, bringing opposable "lips" into close proximity and
forming a cylinder around the duplex (13). Further crystal structures suggest that this
cylinder must be flexible to allow rotation of the DNA (14). Interestingly, molecular
dynamics simulations suggest that the protein accommodates this rotation differently
when it is relaxing positive, rather than negative, supercoils. When the enzyme is
releasing positive supercoils, the "lips" open slightly, while the "hinge" side opens up to
release negative supercoils (15). Since relieving supercoils of opposite signs requires
rotation of the nicked strand in opposite directions, these structural adjustments differ in
order to accommodate the sign of supercoiling being relieved.
While human topoisomerase I is one of the better-characterized proteins that
shows differential interaction, in theory any enzyme that interacts with DNA directly
could be affected by supercoiling. One such class of enzymes is comprised of the Type II
restriction endonucleases, a diverse group that cleaves DNA in a site-specific fashion.
Restriction enzymes are classified as Type II on the basis of their production of
reproducible fragmentation patterns and cleavage at fixed sites at or near their recognition
sequence (16). This broad definition allows for significant diversity within the type and
several subdivisions have been created to classify enzymes based on such factors as
140
subunit composition, nature of recognition site, number of targets, and associated
methylation activity (17). The utility of Type II enzymes as tools for molecular biology
has led to a widespread search for new enzymes and to date more than 3,500 have been
identified (18). Because so many enzymes have been identified in a relatively short
period of time, however, the details of target site recognition and reaction mechanism for
many of these enzymes remain unknown.
Evidence suggests that negative supercoiling may interfere with the ability of
some Type II enzymes to cleave DNA; for some enzymes, supercoiled plasmid with a
single recognition site is not cut as efficiently as linear , DNA (19). An exact role for
supercoiling, however, isn't entirely clear, as other factors can affect cleavage as well.
These include requirements for an allosteric effector, or interaction with bases outside of
the enzyme's recognition sequence.
In this study, I have tested three restriction enzymes (NarI, Ehel, and EcoRI) with
relaxed and positively and negatively supercoiled substrates in order to investigate their
sensitivity to supercoiling. Only NarI and EcoRI showed subtle changes in reactivity
with the introduction of supercoiling. While none of the enzymes tested showed
substantial changes, this class of enzymes is sufficiently diverse that it remains possible
that some may be strongly affected. Identifying such enzymes for further study would
provide valuable information about the ways in which supercoiling affects interactions
between DNA and its modifying enzymes.
141
MATERIALS AND METHODS
Preparationofsupercoiledsubstrates. Positively supercoiled pUC18 substrates
were prepared as described in Chapter 2. Relaxed and negatively supercoiled substrates
were prepared in the same manner as the positive, substituting heat-inactivated chicken
blood extract (CBE) for live enzyme at the appropriate steps to obtain substrates with the
desired presence and direction of supercoiling. All substrates were dialyzed once in 2 L
of TE buffer with 3 M NaC1, using 100,000 MWCO dialysis tubing, followed by four
changes of 2 L of TE buffer, prior to use in enzymatic experiments.
Enzymes and enzyme reactions. Restriction enzymes NarI and EcoRI were
obtained from New England Biolabs (Ipswich, MA). EheI was obtained from Fermentas
(Hanover, MD). All reactions took place in the appropriate lx buffer conditions, using
the buffer provided by the manufacturer. These conditions were as follows. NarI: 10
mM bis-Tris-propane-HC1, 10 mM MgCl 2, 1 mM dithiothreitol, pH 7.0; EcoRI: 100 mM
Tris-HC1, 50 mM NaC1, 10 mM MgCl 2, 0.025% Triton X-100, pH 7.5; Ehel; 33 mM
Tris-acetate, 10 mM magnesium acetate, 66 mM potassium acetate, 0.1 mg/mL BSA, pH
7.9.
Enzymatic reactions were carried out at 37 TC in 200 [L total volume of the
appropriate buffer, with 5 nM DNA concentration. Reactions were initiated by addition
of 5 ýtL of the appropriate diluted enzyme, at concentrations of 40 U/mL for NarI, 10
U/mL for EcoRI, and 5 U/mL for Ehel. At pre-determined time points, 10 [tL aliquots
were removed from the reaction and quenched by addition of 10 1 iL of nuclease stop
142
solution (100 mM EDTA, 100 mM Tris, pH 8.0, 40% sucrose, 0.1% bromophenol blue).
Samples were resolved at 4.5 V/cm for 90 minutes on 1% agarose gels in lx TBE
containing 0.5 [tg/mL ethidium bromide. Relative fractions of open circular, linear, and
closed-circular plasmid were quantified based on ethidium fluorescence, as described in
Chapter 3.
RESULTS
The effects of supercoilingon the activity of restrictionenzymes. Several factors
can affect cleavage of cognate DNA by restriction enzymes, including buffer conditions,
sequence flanking the recognition site, and the number of recognition sites present on the
molecule. By using plasmids that differ only in supercoiling status, and carefully
controlling reaction conditions, other factors can be ruled out and any changes in
reactivity can. be attributed to supercoiling.
Type II restriction enzymes cleave DNA at a fixed site at or near their recognition
sequence. Cleavage of a molecule to linear form can result from one of two modes (Fig.
6.1) (20-22). In the first, intact DNA is bound by the enzyme, which cleaves both strands
of the DNA in a single binding step. In the second, the enzyme cleaves a single strand
before releasing a nicked molecule, and the intact strand is later cleaved in a second
binding event. Since our relaxed and supercoiled substrates are all closed-circular
molecules, the products of reaction with restriction enzymes offer insight into these
cleavage modes. Conversion of supercoiled plasmid exclusively to linear form indicates
the first, two-strand cleavage mode is occurring. Conversely, appearance of open circular
143
(nicked) plasmid in the reaction products is indicative of the second mode. In the second
mode, linear products will also appear as the open circular molecules are cleaved. Any
given enzyme does not necessarily follow a single reaction pathway; in fact, some
enzymes have been shown to partition between the two under some circumstances (22).
NarIcleaves both strandsslightly more efficiently in relaxedplasmid DNA. NarI
is a type II restriction enzyme that cleaves at the palindromic sequence 5'-GG CGCC'3to leave a two base 5' overhang (23). It has been reported to cleave supercoiled pUC19
20 times less efficiently than linear X DNA (19). The enzyme cleaves considerably more
efficiently when a second recognition site is present (24), but this requirement for an
allosteric effector likely would not explain the reported difference in cleavage between
supercoiled pUC 19 and linear X, as both contain a single NarI recognition site. NarI
cleavage may be also be affected by flanking sequence, cleaving more efficiently in
certain contexts. Such sequence differences could contribute significantly to the
observed cleavage variations, although supercoiling may also play a role.
To ascertain the effect of supercoiling on cleavage by NarI, the enzyme was
reacted with relaxed, positively and negatively supercoiled DNA and the kinetics of
reactions were monitored. As shown in Figure 6.2, the overall rate of cleavage was not
significantly different among the different states of supercoiling. Overall, the plasmids
were cleaved relatively slowly, which is expected for a single-site substrate with an
enzyme that requires an allosteric effector. Closed-circular plasmid disappeared at
approximately the same rate for all three forms. A difference arises, however, when the
rate of conversion to linear and to open circular forms are analyzed separately. While all
144
three forms are predominately converted to open circular, a small fraction of relaxed
plasmid is converted to linear.
NarI has previously been demonstrated to cleave negatively supercoiled plasmid
via the two-binding-event pathway, cleaving one strand of the substrate during an initial
binding and returning to convert open circular plasmid to linear at a much slower rate
(21). This pathway is followed regardless of whether or not an allosteric effector is
present (21). The small amount of linear plasmid formed from relaxed plasmid, however,
is likely due to a partitioning between the two modes of cleavage. Open circular plasmid
is structurally identical, regardless of the original supercoiled state of the molecule, so it
would not be converted to the linear form faster in one reaction than in the others. The
formation of linear plasmid should therefore be the result of cleavage of both strands in a
single binding event.
EheI is not sensitive to eitherpositive or negative supercoiling. Ehel is a
neoschizomer of NarI that leaves a blunt end at 5'-GGC GCC-3' sites (25). The enzyme
does not require an allosteric effector, and has been shown to cleave both strands of the
recognition sequence in a single binding step (21). As can be seen in Figure 6.3, Ehel is
largely insensitive to supercoiling, cleaving at approximately equal rates in relaxed, and
positively and negatively supercoiled plasmid. Supercoiling also did not affect the
cleavage mode of the enzyme, with direct conversion to linear seen in all cases.
Positive supercoilingslightly increases the rate of cleavage by EcoRI. EcoRI is
perhaps the best-studied of all restriction enzymes, cleaving at G AATTC sequences to
leave a four base 5' overhang (26). The crystal structure of EcoRI bound to its
recognition site has been solved, and the protein is shown to distort the DNA upon
145
binding. EcoRI induces a kink in the DNA backbone that is accompanied by widening of
both grooves, an increased rise between base pairs and a slight unwinding of the helix
(27). The enzyme can cut either one or both strands in a single binding event (20); the
partitioning between single and double cleavage in the first binding event appears to be at
least partially determined by the sequence flanking the recognition site (22). EcoRI is
reported to cut X DNA somewhat more efficiently than it does supercoiled plasmids
pUC19 and pBR322 (19), although the reason for this differential cleavage is not known.
In these studies, EcoRI predominately cleaved directly to linear for all three
forms, producing only a small amount of open circular plasmid (Fig. 6.4). The enzyme
was slightly affected by supercoiling, cleaving positively supercoiled plasmid at a faster
rate than either relaxed or negatively supercoiled plasmid. It is not clear if the change in
reactivity is accompanied by a change in partitioning between one- and two-strand
cleavage in the first binding event; both the linear and open circular fractions are
increased with positive supercoiling relative to the other two forms.
DISCUSSION
Type II restriction enzymes comprise a broad class that act to cleave DNA at a
fixed site at or near their recognition sequence. Previous studies have suggested that
several of these enzymes may be sensitive to DNA supercoiling, although many of these
reports are in conflict (19,28-3 0).
In this study, I have shown that cleavage by restriction enzymes can be affected
by supercoiling, although the differences observed were small. Supercoiling could
146
conceivably affect the interaction of these enzymes with DNA in a number of ways,
including altering binding of the enzyme and product release.
Specific binding of several Type II enzymes is known to induce conformational
changes in the DNA structure, including kinks or bends in the DNA backbone, and
unwinding of the helix (27,31). Supercoiling-induced structural changes may affect the
ease with which an enzyme can induce these changes. A negatively supercoiled
molecule, for example, is more readily unwound than a positive one, since unwinding is
already present in the negative molecule. Additionally, supercoiling-induced changes in
the structure of the DNA grooves could alter how the enzyme is able to track along the
helix during facilitated diffusion, potentially affecting location of the target site.
Supercoiling could also contribute to changes the rate at which the enzymes
release cleaved product. Product release is the rate-limiting step for several Type II
enzymes under certain conditions (30). When the backbone of a supercoiled molecule is
cleaved, the free energy of supercoiling that is trapped in the molecule is held in place
only by the bound enzyme. The force of this elastic torsion may be sufficient to drive the
enzyme free from the target site at an accelerated rate, allowing for a faster overall
cleavage rate by the enzyme. It also may manifest by changing the partitioning between
one- and two-strand cleavage in the first binding event. Such a change in partitioning
might explain the small fraction of relaxed plasmid that NarI is able to cleave directly to
linear; the lack of supercoiling in the molecule may allow the enzyme to remain bound
after the first cleavage event in some fraction of the molecules.
The interaction of restriction enzymes with DNA is a complex one, which could
be affected by supercoiling in many different ways. Because of this complexity, it is
147
impossible to attribute any difference in reactivity to a given cause without more detailed
study. The changes in reactivity I have shown here are small, and provide limited
incentive for further investigation. Type II restriction enzymes as a class, however, are
sufficiently diverse that it is reasonable to expect an enzyme to be identified with more
pronounced supercoiling sensitivity. Once such an enzyme was located, more in depth
studies could be employed to elucidate the specific changes supercoiling produces.
148
H
H
B
C
Figure 6.1. Possible reaction modes of closed circular DNA with restriction enzymes. A)
Relaxed, negatively and positively supercoiled plasmid are all covalently closed circular
molecules. B) A restriction enzyme binds at its recognition site, and cleaves both strands of
the target DNA in a single binding event. C) Alternatively, the restriction enzyme might bind
at its recognition site, and cleave one strand in the first binding event. This leaves an open
circular molecule. The enzyme then returns to cleave the molecule to linear in a second
binding step.
149
*
4
A
Positive
Negative
Relaxed
~-
I
14
~----~-`---"-11-"11`I----~-"~
I
15
10
4
0
100
4
. ........
.
Time (minutes)
Figure 6.2. Kinetics of reaction of NarI with supercoiled and relaxed substrates. Relaxed
plasmid is cleaved by NarI endonuclease directly to linear at a slightly faster rate than either
positively or negatively supercoiled plasmid. Plasmid substrates were reacted with NarI as
described in Materials and Methods. Data points shown are the average of four replicates; error
bars represent the standard error.
150
S
L
~~~1
0 Negative
A. Positive
A Relaxed
.......
.....
0
0L
Ol
ý-
I
0.
o
I
I
4
I
1
"iIts* a r. I
A;
t
RI~irr
7----1--
-----
dTh
-t----
;------------------------·-----------------....
...
.......
..
......
.
10
f
0____
i
_
_J
ioo100
I,
75
£
I
50
25
Time (minutes)
Figure 6.3. Kinetics of reaction of Ehel with supercoiled and relaxed substrates. Cleavage
progresses directly to linear; rates are the same regardless of supercoiling state. Plasmid
substrates were reacted with Ehel as described in Materials and Methods. Data points shown are
the average of four replicates; error bars represent the standard error.
151
20
15
0
100
(
---------
I
~-LIIIII^·l~·--·-·-··---·--~
I
75
50
25
20
40
60
Time (minutes)
Figure 6.4. Kinetics of reaction of EcoRI with supercoiled and relaxed substrates. Positively
supercoiled plasmid is cleaved by EcoRI endonuclease at a slightly faster rate than either relaxed
or negatively supercoiled plasmid. Plasmid substrates were reacted with EcoRI as described in
Materials and Methods. Data points shown are the average of four replicates; error bars
represent the standard error.
152
LITERATURE CITATIONS
1.
Wu, H.Y., Shyy, S.H., Wang, J.C. and Liu, L.F. (1988) Transcription generates
positively and negatively supercoiled domains in the template. Cell, 53, 433-440.
2.
Giaever, G.N. and Wang, J.C. (1988) Supercoiling of intracellular DNA can occur
in eukaryotic cells. Cell, 55, 849-856.
3.
Ljungman, M. and Hanawalt, P.C. (1992) Localized torsional tension in the DNA
of human cells. Proc Natl Acad Sci USA, 89, 6055-6059.
4.
Steck, T.R., Franco, R.J., Wang, J.Y. and Drlica, K. (1993) Topoisomerase
mutations affect the relative abundance of many Escherichia coli proteins. Mol
Microbiol, 10, 473-481.
5.
Vologodskii, A.V., Levene, S.D., Klenin, K.V., Frank-Kamenetskii, M. and
Cozzarelli, N.R. (1992) Conformational and thermodynamic properties of
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Chapter 7 - Conclusions
In the previous chapters, I described the preparation of highly purified supercoiled
substrates and how I used these substrates to demonstrate that physiological levels of
supercoiling induce subtle changes into the structure of DNA. This supercoiling can also
affect the interaction of DNA with various DNA-modifying enzymes, as can be seen by
the small changes that I have shown in cleavage of DNA by Type II restriction enzymes.
A rigorous method for the preparation of supercoiled DNA substrates was crucial
to these studies with the inherent difficulty in preparing highly positively supercoiled
plasmid serving as a hindrance in previous studies of the phenomenon. On the basis of a
method developed earlier in this lab (1), I undertook the development of a new method
for producing highly positively supercoiled substrates. This was necessary due to
variable topoisomer distributions, high degree of nicking, and batch-to-batch variability
in reactivity of substrates prepared using the original method
The new method employs more stringent quality controls in order to ensure high
levels of positive supercoiling, to remove differences between substrates that are not due
to supercoiling, and to reduce the population of open circular molecules that contaminate
the preparations. Additionally, I have expanded this method to allow the use of a simple
salt supplement to control the direction of supercoiling that is induced. This new method
allows generation of either positively or negatively supercoiled substrates, and should
facilitate studies of supercoiling in which the desired substrate cannot be propagated in a
bacterial system, such as DNA circles with non-canonical bases or base adducts inserted.
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To assess the solvent accessibility of supercoiled substrates, relaxed and
positively and negatively supercoiled plasmids were reacted with both enzymatic and
chemical probes of DNA structure. Whereas negatively supercoiled plasmid was very
sensitive to cleavage by the single strand-specific nucleases S1 and BAL-31, no increased
reactivity was seen with positively supercoiled substrates. Chloroacetaldehyde, which
reacts with the hydrogen bonding region of the DNA bases to form cyclic etheno adducts,
did not show an increase in either 1,N6-ethenoadenine or 3,N4-ethenocytosine with either
negative or positive supercoiling. The result with negative supercoiling is somewhat
surprising, since negative supercoiling is known to induce denaturation of the DNA helix
with increased solvent exposure of the bases for reaction with chloroacetaldehyde. Still,
the nature of the denaturation in negatively supercoiled plasmid may be transient, which
might prevent etheno adduct formation from occurring at an increased rate.
Chloroacetaldehyde forms etheno adducts in a multi-step reaction process and only the
end products are being analyzed in our system. It is possible that the rate-limiting step of
the chloroacetaldehyde reaction is slower than the base pair opening rates in the various
supercoiled substrates.
The reactivity of negatively supercoiled plasmid that is seen here is consistent
with a molecule that has transient denaturation of the DNA strands, but without any
stable supercoiling-induced secondary structures. The results with positively supercoiled
plasmid are less informative. The lack of reactivity with single strand-specific nucleases
could indicate a lack of increased solvent exposure of the bases, although alternative
possibilities remain. The structure that is recognized as a target for cleavage by these
enzymes is not clear. It may be possible that the structure of positively supercoiled
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plasmid is changed in such a way that the enzyme is unable to identify it as a cleavage
target. Another possibility is that the rate at which the bases are solvent exposed with
positive supercoiling is kinetically too fast to be recognized by the enzymes. The lack of
reactivity with chloroacetaldehyde is relatively uninformative, since no increase is seen in
reactivity with negatively supercoiled plasmid with its known denaturation. The
strongest conclusion that can be drawn from this data is that an alternative secondary
structure with stably unpaired bases is not forming in the positively supercoiled
substrates.
Studies with other structural probes may prove helpful in furthering our
understanding of structural changes induced by positive supercoiling. While the rate of
formation of the EA and EC adducts may not be a useful measure of solvent exposure, it
has been suggested that the partitioning between the two ethenoguanine adducts (1,N2ethenoguanine and N2 ,3-ethenoguanine) can be affected by base-pairing (2). The N1
position is blocked by hydrogen bonding, so partitioning would shift to favor the N2 ,3EG
adduct in the case of base pairing. Comparing the relative levels of these two adducts in
our substrates may prove more informative than studying the absolute levels of EA and
EC. Dimethyl sulfate should also be investigated further. Preliminary studies in this lab
have suggested that positively supercoiled DNA is methylated at an increased rate at both
GN 7 and AN3 by dimethyl sulfate, relative to negatively supercoiled plasmid.
Unfortunately, studies on relaxed substrate have not been performed. Since the reactivity
of relaxed substrates provides a crucial baseline for evaluating the level of reactivity that
is not caused by supercoiling, the lack of this data prevents detailed interpretation of
these results. Expanding the dimethyl sulfate studies to include relaxed substrates should
159
provide useful information about the status of the GN 7 and AN 3 sites in supercoiled
substrates. Reactivity with other chemical probes of DNA structure, such as potassium
permanganate and diethyl pyrocarbonate, should also be considered.
To examine the fine structural changes that occur as a result of positive
supercoiling, we employed Raman difference spectroscopy. The information-dense and
well-defined Raman signature of DNA is very sensitive to conformational changes in the
molecule, and by digitally subtracting the spectrum of relaxed plasmid from that of a
supercoiled substrate, we can identify supercoiling-induced changes in structure. All
forms were shown to be in predominately B-form conformation, with no identifiable
conversion to alternative structures such as Z-DNA occurring. This is consistent with our
chemical and enzymatic data.
The structural changes that did occur with supercoiling were reproducible, but
subtle, suggesting that at the levels of supercoiling tested, the molecules accommodate
supercoiling by making small structural adjustments. These adjustments were largely in
the sugar-phosphate backbone of the DNA, with a smaller contribution from a slight
disruption of the base environment. Importantly, the difference spectra showed that
while some changes occurred in both forms of supercoiling, others were unique to the
direction of supercoiling induced. In negatively supercoiled plasmid, the most notable of
these was a difference peak diagnostic of a change in the torsion angles of the phosphate
backbone. For the positive, a series of difference peaks suggests that a small fraction of
sugars in the C3' endo form is converting to a C2' endo form that is more consistent with
B-DNA. Since conversion from the C3'endo to the C2'endo sugar pucker allows
adjacent phosphates to be farther apart by -1 A, this makes sense in the context of the
160
lengthening of the backbone that is required with positive supercoiling. It is worth noting
that the formation of "P-DNA" at high levels of introduced twist is thought to result from
the phosphate backbone having reached maximal bond lengths, and then adapting to
additional twist by moving to the center of the helix to reduce the helical radius (3). The
change in sugar conformation seen in my experiments may then be an early step in this
process.
Other notable changes that are seen in the positively supercoiled substrates
include an indication of altered hydrogen bonding at the GN 7 position, and a
destabilization of base pairing. Located in the major groove, the GN 7 position may be
experiencing a disruption in solvent interactions as a result of the bending and twisting
motions of supercoiled DNA. Free energy studies have determined that the entropy of
supercoiling is large and positive. Since supercoiling limits the structures that a molecule
can sample, the configurational entropy would be expected to be negative and another
entropy component would have to be responsible for the positive change. This increased
entropy has been proposed to result from the disruption of the interaction of DNA with
solvent and other molecules, due to the motions of the supercoiled DNA (4). This would
be consistent with our observations at GN 7. Further evidence of this phenomenon could
be obtained from the results of the chemical studies discussed previously, as the GN 7 is a
target site for dimethyl sulfate methylation. Two peak shifts in the positively supercoiled
difference spectra suggest destabilization of base pairing. Taken together with enzymatic
studies, this suggests that while stable eversion of the bases from the helix is not
occurring, some degree of base pair disruption is caused by physiological levels of
positive supercoiling.
161
In order to examine the effects of supercoiling at a wide range of superhelical
densities, I designed and began preparation of 2-aminopurine-containing DNA substrates
for use in a magnetic micromanipulator. In such a system, a long segment of DNA would
be anchored on one end to a solid support, and on the other end to a magnetic bead which
could be used to introduce precise amounts of twist into the molecule. By incorporating a
base analog with known fluorescence properties, changes in the DNA structure could be
monitored fluorimetrically. Unfortunately, due to instrument sensitivity, the experiments
are not technically feasible. The design of the substrate has been optimized in such a way
to significantly improve any future studies. The time and effort involved in preparing
these substrates is substantial, however, so it is crucial to verify adequate instrument
sensitivity with the proposed system before proceeding with substrate construction.
I then proceeded to investigate the effects of supercoiling on interaction between
DNA and the structurally-sensitive Type II restriction enzymes. In two of the three
enzymes tested, a slight sensitivity to supercoiling was seen. The interaction between
these enzymes and DNA is complex, however, and particularly with such small
differences, the mechanisms behind the observed changes could not be attributed.
Screening of these enzymes is fairly easy, however, and with the diversity of Type II
enzymes it seems reasonable that one might be identified with more substantial
supercoiling-induced changes in reactivity. A more detailed study of such an enzyme
should provide important new information about the effects of supercoiling on proteinDNA interactions.
Overall, I have demonstrated that positive supercoiling has subtle, but distinct
effects on the structure of DNA, which are consistent with the mechanical effects of
162
overwinding. With the ubiquity of supercoiling in cells, and the structural sensitivity of
many classes of DNA-interactive enzymes, it is likely that even these small changes can
have significant physiological effects. Further study of both the structural consequences
of positive supercoiling, and the effects that this has on enzymes that interact with DNA,
should help to clarify the biological role of this important phenomenon.
163
LITERATURE CITATIONS
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Guengerich, F.P. and Persmark, M. (1994) Mechanism of formation of
ethenoguanine adducts from 2-haloacetaldehydes: 13C-labeling patterns with 2bromoacetaldehyde. Chem Res Toxicol, 7, 205-208.
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Allemand, J.F., Bensimon, D., Lavery, R. and Croquette, V. (1998) Stretched and
overwound DNA forms a Pauling-like structure with exposed bases. Proc Natl
AcadSci USA, 95, 14152-14157.
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Vologodskii, A.V. and Cozzarelli, N.R. (1994) Conformational and
thermodynamic properties of supercoiled DNA. Annu Rev Biophys Biomol Struct,
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164