THE CATHOLIC UNIVERSITY OF AMERICA
Studies of Single DNA-Histone Binding Events Using a Novel Magnetic Tweezer
A DISSERTATION
Submitted to the Faculty of the
Department of Physics
School of Arts and Sciences
Of The Catholic University of America
In Partial Fulfillment of the Requirements
For the Degree
Doctor of Philosophy
By
Christopher P. McAndrew
Washington. D.C.
2012
Studies of Single DNA-Histone Binding Events Using a Novel Magnetic Tweezer
Christopher McAndrew Ph.D.
Director: Abhijit Sarkar, Ph.D.
How do proteins, specifically histones, react to low levels of applied tension?
Until now, low-force, single-molecule studies of DNA-protein interactions have been
hampered by experimental limitations in the devices used to observe protein binding and
unbinding events. To overcome this, I report on the development of a new magnetic force
transducer or “tweezer” that can apply pico-Newton forces on single DNA molecules in
the focus or horizontal plane and allow for the observation and measurement of single
DNA-protein interaction events. This semi-closed, horizontal magnetic tweezer allows us
to gather high resolution data on changes in DNA’s end-to-end extension because these
changes are coplanar with the pulling force. DNA constructs (lambda-DNA end labeled
with a 3µm polystyrene bead and a 2.8µm paramagnetic sphere) and appropriate buffer
(1X Tris EDTA) are introduced to a small volume semi-closed cell created using two thin
cover-slips and 1mm glass spacers. This semi-closed configuration isolates a sample and
produces low-noise force and extension measurements. With this new instrument, I report
on the successful study of nucleosomal kinetics and energetics. Observation of individual
nucleosome ruptures at and above a critical force of 1.6pN is first reported here. I claim
that the capability of the instrument to produce low force-loading rates on DNA with
bound proteins is responsible for this observation. This capability also allowed me to
make a comparative study of nucleosomes formed using both native (typical to normal
physiological conditions) and post-translationally – hyper-acetylated - histones. The
experimental technique developed allowed for the observation of unbinding events at
low-forces and the subsequent calculation of the differences in the binding affinities of
both
native
histones
and
post-translationally
modified
histones.
This dissertation by Christopher McAndrew fulfills the dissertation requirement for the
doctoral degree in Physics approved by Abhijit Sarkar, Ph.D., as Director, and by
Frederick Bruhweiler, Ph.D., and Franz Klein, Ph.D., and Pamela Tuma, Ph.D. and
Patrick Mehl, Ph.D., as Readers.
Abhijit Sarkar, Ph.D., Director
Frederick Bruhweiler, Ph.D., Reader
Franz Klein, Ph.D., Reader
Pamela Tuma, Ph.D., Reader
Patrick M. Mehl, Ph.D., Reader
ii
Dedication
I dedicate this thesis work to my ever patient and understanding parents:
Charles and Mary Ann McAndrew
iii
Table of Contents
Chapter 1
Introduction
1.1 DNA
1.2 Component Histones
1.3 Nucleosomes
1.4 Histone Post-Translational Modifications
1.5 Response of Single DNA Polymer to Micromechanical Manipulation
1.6 Response of Single DNA-Protein Complexes to Micromechanical Manipulation
1.7 Response of Chromatin, Nucleosomes and Nucleosomal Arrays to Micromechanical
Manipulation
1.8 Experimental Methods for single molecule micromanipulation
1.8.1 Optical Tweezers
1.8.2 Atomic Force Microscopy (AFM)
1.8.3 Vertical Magnetic Tweezers
1.8.4 Transverse or Horizontal Magnetic Tweezers
1.9 Research Problem and Approach
1.10 Results
1.11 Plan of the work
Chapter 2
Horizontal Magnetic Tweezer
iv
2.1 Description of system
2.2 Preparation of DNA, Beads, Buffer Chamber, Pipettes, Buffers, Protein Solutions
2.3 Data Analysis
2.3.1 DNA Tether Extension Measurement
2.3.2 Force Measurement by Fluctuation-dissipation theorem
2.3.3 Measurement of Bead velocities
2.3.4 Detection of Steps
2.4 Calibration
2.4.1 Extension Calibrations-Graticule
2.4.2 Force Calibration - Stokes Law Measurement
2.4.3 Fit to WLC with correct persistence length
2.5 Fit to WLC with correct persistence length
2.6 Typical Low-Force-Extension Experiment for DNA
Chapter 3
Pulling on Nucleosomes
3.1 Instrumentation & Technique
3.1.1 Introductory Material
3.1.1a Magnetic Tweezers
3.1.1b Histone purification
3.1.1c Histone experiments
3.2 Results for Pulling on Nucleosomal Arrays
v
3.2.1 Nucleosome Array formation and force-induced dissociation
3.2.2 Nucleosomal Arrays Under Tension
3.2.3 Step Size Distribution
3.2.4 Nucleosome Response to Force: Step-Counts and Step-Size
3.3 Discussion
3.3.1 Advantages of this System Over Other Systems
3.3.2 Low Force Response: Theoretically-Predicted Critical Force Observed
3.3.3 Moderate and High Force Response
3.3.4 Free Energy vs. Force
3.3.5 Step-Size and Applied Force
3.3.5 Step Size Does Not Depend on Force
Chapter 4
Native vs. Post-Translationally Modified Histones
4.1 Post-Translational-Modifications: Structure and Significance
4.2 Comparison Study of Native and Post-Translationally Modified Histones
4.2.1 Protein Introduction & Re-extension
4.2.2 Step Histogram
4.2.3 Force Histogram & Force-Step hybrid Histogram
4.2.4 Step-Duration Histogram
4.2.5 Free-Energy Comparison
4.3 Discussion
vi
4.3.1 Comparison to Other Works
4.3.2 Observations on the Results
Chapter 5
Conclusion
5.1 Summary of Results
5.2 Future Directions
Appendices
1) Appendix I: Tris-EDTA Buffer Preparation
2) Appendix II: DNA End-Functionalization Protocol
3) Appendix III: Labeling of 3µm diameter polystyrene beads with anti-digoxygenin
4) Appendix IV: Micro-injection protocol
5) Appendix V: Protocol for creating and using micropipettes for aspiration
6) Appendix VI: Native Histone Preparation
7) Appendix VII: Hyper-Acetylated Histone Preparation
8) Appendix VIII: Derivation of the Fluctuation-Dissipation Theorem
9) Appendix IX: Sterilization and cleaning protocols for all Instrument
10) Appendix X: Preparation and Construction of Buffer Chamber
11) Appendix XI: DI water filtering and sterilization
12) Appendix XII: Free Energy Calculation
vii
List of Figures
Chapter 1
1) Figure 1.1: DNA structure and physical characteristics
2) Figure 1.2: Nucleosome: Core Histones and DNA
3) Figure 1.3: DNA Force-Extension Curve
4) Figure 1.4: Laser Tweezers Setup, Dual Trap
5) Figure 1.5: Vertical Magnetic Tweezer Schematic
Chapter 2
1) Figure 2.1: Physical Principle of Transverse Magnetic Tweezer
2) Figure 2.2: Our Horizontal Magnetic Tweezer and Buffer Chamber
3) Figure 2.3: Force vs. Distance Curve – Calibration and Experiment Results
4) Figure 2.4: Force-Extension Curve in Force Range of 1pN - 20pN
5) Figure 2.5: Hysteresis Curve for DNA
Chapter 3
1) Figure 3.1: Image of Gel Highlighting Histones
2) Figure 3.2: Time Trace of a Typical DNA-Histone Experiment
3) Figure 3.3: Sketch of Experimental Events
4) Figure 3.4: Low-Force Response of Nucleosomes
5) Figure 3.5: Moderate-Force Response of Nucleosomes
6) Figure 3.6: High-Force Response of Nucleosomes
7) Figure 3.7: Nucleosome Step-Size Histogram
8) Figure 3.8: Nucleosome Force-Event Histogram
viii
9) Figure 3.9: Nucleosome Step-Size and Step-Force Histogram
Chapter 4
1) Figure 4.1: Principle of the Experiment
2) Figure 4.2: A and B: Typical Time-Trace for Native and PTM experiments
3) Figure 4.3: A and B: Histograms of Step-Sizes for Native and PTM Nucleosomes
4) Figure 4.4: A and B: Histograms of Disruption Event-Forces for Native and PTM
Nucleosomes
5) Figure 4.5: A and B: Histograms of Disruption Step-Sizes and Event-Forces for
Native and PTM Nucleosomes
6) Figure 4.6: Average and Median Time Between Nucleosomal Disruptions for
Native and PTM Nucleosomes
7) Figure 4.7: Average and Median Free-Energy Comparison of Native and PTM
Nucleosomes
ix
Acknowledgments
I gratefully acknowledge the assistance of the following individuals: Christopher Tyson,
Joseph Zischkau, Ryan Vilbig, Pamela L. Tuma, Patrick Mehl, Ian Pegg, Robert Mohr,
Benjamin Buckeye, Helen DeCelles-Zwerneman, Jonathan Luke, Eric Fischer, Paul
Branch, Venigalla Rao, Mark Knepper, C.L. Chou, & Dunja Skoko. Funding from
Vitreous State Laboratory & The Catholic University of America is gratefully
acknowledged.
x
Chapter 1
Introduction
1.1 DNA
DNA is the storage system for the genetic information of all living things. As the
carrier of genetic information, DNA is responsible for 1) passing on hereditary traits from
one generation to the next, 2) maintaining the blueprints for all of the proteins needed
within the cell, and 3) synchronizing the formation of those proteins within the cell with
specific events in the cell cycle.
DNA is a long, semi-flexible, charged polymer – a polymer is a molecule made of
many repeat monomer subunits - chain subject to physical laws. DNA in the nucleus
must be compacted within the small nuclear volume, and at the same time the
information stored in the long DNA chain must be accessible, easily read and efficiently
repackaged without loss or damage of stored data.
Friedrich Miescher discovered a substance in cellular nuclei that was rich in
phosphorous and nitrogen and called it nuclein (Dahm, 2005). Albrecht Kossel then
discovered that nuclein was composed of four bases, sugar molecules, and phosphate
groups (Kossel, 2011). Avery, Mcleod, and McCarthy in 1944 realized that DNA was the
carrier of genetic information (Avery, 1944). They designed an experiment in which
DNA from one strain of common bacteria was transferred to another strain of the same
bacteria. Upon transfer, it was observed that the host bacteria took on the physical
characteristics of the donor. Then, in 1950, Erwinn Chargaff found that the nucleotide
1
2
monomers adenine (A), thymine (T), cytosine (C), and guanine (G) that composed DNA
were found in specific ratios (Chargaff, 1950). The ratios of adenine to thymine and of
cytosine to guanine were found to be 1:1. Subsequent X-ray diffraction experiments
provided evidence of a basic repeatable structure. Other experiments found that there
exists a strong affinity for A to pair with T, and likewise for C to pair with G.
The groundwork was laid for Francis H.C. Crick and James D. Watson to piece
together a coherent description of the structure and function of DNA. They proposed the
double-helical structure (Crick and Watson, 1953) known today based on DNA X-ray
diffraction data obtained by Rosalind Franklin and Maurice Wilkins (Wilkins et al,
1953). The basic monomer repeat units A, T, C, and G formed individual long
polynucleotide chains held together by covalent bonds in a sugar-phosphate backbone.
Two such chains with complimentary sequences of A, T, C, and G could pair with the
bases pointing inward and facing each other, and form an intertwining right-handed
double helical structure that was extremely stable.
In 1961, Crick et al performed experiments that showed that a sequence of three
bases coded for a single amino acid (1961). This finally established how genetic
information is coded in DNA.
The structure of DNA is now understood: DNA is a right-handed double helix
formed through the complimentary base-pairing of two polynucleotide chains. While
each strand of DNA is a covalently-linked polymer of monomers corresponding to A, T,
G, and Cs, the double helix itself is stabilized by hydrogen bonds contributed by the
complimentary pairing of bases A with T and G with C and base stacking interactions
3
which are entropic in origin.
The monomer building blocks of DNA are nucleotides. The nucleotides are
composed of a five carbon sugar ring called deoxyribose, a phosphate group, and one of
four bases: adenine (A), thymine (T), cytosine (C), and guanine (G). The carbons in the
sugar are labeled 1’ – 5’, with the phosphate group attached at the 5’ site and capable of
forming a phosphodiester bond with a 3’ carbon on a neighboring sugar. Nucleotide
chains are thus linked together through covalent bonds between the 3’ carbon of one
nucleotide and the 5’ carbon of another nucleotide thus forming the sugar phosphate
backbone, which gives the chain directionality. The two strands that form DNA are
aligned anti-parallel to each other, so that at both ends of linear DNA a free 3’ end on one
strand is associated with a free 5’ end on the other strand. See figure 1.
When two complimentary polynucleotide chains come together, hydrogen bonds
form between the A on one strand and the T on the other, and between C on one strand
and the G on the other. Specifically, three hydrogen bonds form between C and G; two
hydrogen bonds form between A and T. A and T are referred to as a complimentary bases
as are G and C. Thus, two polynucleotide chains with fully complimentary sequences
considered in the opposite 3’-5’ and 5’-3’ senses can entwine together to form the double
helix (Darnell et al, 1986).
The sequence in which the nucleotides are connected determines the primary
structure of DNA. The spontaneous double helical configuration of the two strands is
referred to as the secondary structure of DNA. A long chain like DNA in the nucleus can
also have higher levels of spatial organization referred to as the tertiary structure of DNA.
4
Polymers are molecules composed of many repeating subunits called monomers.
If all of the subunits are the same, the polymer is referred to as a homopolymer. If there is
more than one type of monomer, it is called a hetero-polymer. Polymers can be stiff,
flexible or semi-flexible; they can be of various lengths and topologies: linear, ring
shaped, or branched; they can be charged or neutral (charged polymers are called
polyelectrolytes).
DNA is one of the longest polymers in nature and is far longer than any known
synthetic polymer - it can vary from tens of microns for viral DNA to several tens of
centimeters for DNA in eukaryotic cell nuclei. DNA is semi-flexible which means it is
highly flexible on long length scales, but it is nearly rigid for lengths less than ~50nm,
this 50nm is referred to as the persistence length of DNA (Bustamante et al, 1994), and is
charged with a charge density of two negative charges per base-pair. (However, it should
be noted that the electrostatic interactions between these charges is heavily screened by
salt counter ions.) The corresponding van der Waals diameter is 2nm. Each strand
completes one full rotation around the other every 3.4nm or ~10.5bp (bp = basepair),
which implies that the distance between successive rungs – each rung being a
complimentary base pair - of the DNA is ~0.34nm. This twist rate is also subject to
random thermal fluctuations but is fairly constant over length scales ~100nm or 300 bp.
This length is referred to as the twist rigidity of DNA and is the twist analogue of the
persistence length discussed above. A schematic representation of the double helix and
the hydrogen-bonded nucleotide sequences is shown in figure 1.1.
5
Right Handed Twist
A
T
G
C
Rise
0.34nm
C
G
T
A
Pitch
3.4nm
Hydrodynamic Diameter
2.0nm
Figure 1.1: Here, a simple sketch highlights the fundamental building blocks of DNA and
a few basic physical properties. The figure on the left shows the phosphate group, the
pentose sugar with the carbon atoms labeled 1’ – 5’ and the nucleotide bases adenine,
thymine, cytosine, and guanine. This also shows how the single strands of nucleotide
chains link directionally 5’ – 3’ and 3’ – 5’. The sketch on the right lists several physical
characteristics of the DNA molecule. First, there is 0.34nm between each base pair.
Second, the double helix completes one full twist for every 10.5 base pairs. Third, DNA
in aqueous solution has a diameter of ~2nm. Colored figure courtesy of http://www.ciccaracas.org/departments/science/images/08P-210-DNA-5-3.jpg.
6
1.2 Component Histones
Proteins are polymers formed by stringing together chains of twenty amino acids
in a variety of sequences (Petsko et al, 2004). Each amino acid has a modular structure
consisting of a chemical subunit common to all amino acids. There are subgroups called
residues - of which there are twenty – which give each amino acid its distinct chemical
character. The amino acid sequence of a protein determines its primary structure (Petsko
et al, 2004). A single polypeptide chain or portions of a chain can spontaneously form
two types of secondary structures: -helices and -sheets (Petsko et al, 2004); in general,
both -helices and -sheets can be present simultaneously along different stretches of the
protein. How many -helices and -sheets form and where they form along the molecule
are essentially determined by the underlying amino acid sequence. The -helices and sheets are stabilized by hydrogen and other weak bonds.
The ability of the protein to perform its function in the cell is determined by its
tertiary structure, which refers to a higher level of folding in which the successive
-
helices and -sheets that form along the polypeptide are organized into complex three
dimensional conformations. These conformations are also usually stabilized by weak
hydrogen bond- type interactions, but they are quite robust since many weak bonds
simultaneously contribute to maintaining the spatial organization of the polypeptide chain
(Petsko et al, 2004). Proteins can also form quaternary structures with two or more like or
dissimilar proteins forming stable dimeric or multimeric complexes (Petsko et al, 2004).
7
In fact, many molecular machines involved in basic genetic processes in the cell consist
of just these multimeric structures built out of component proteins.
A particular class of proteins called histones plays a critical role in condensing
DNA so that it can fit inside the nucleus (Darnell et al, 1986). There are five kinds of
histones labeled H1 or H5, H2A, H2B, H3, and H4. Two of each H2A, H2B, H3, and H4
form the eight-protein octomer core. H1 is the linker histone and is believed to add to the
stability of a nucleosome structure by acting as a “seal” tying together the entrance and
exit DNA loop around the core particle. The histones which compose the octomer core
have an N-terminal tail domain which contains positively charged amino acid residues
(Hansen, 2002). This structural conformation, or tail, enhances the histones’ ability to
bind to DNA as will be described below.
1.3 Nucleosomes
The DNA polymer chain of eukaryotes must be confined to a cell nucleus with a
~1-5μm diameter and an approximate volume of ~ 10-16m3. The first physical problem is
how to fit the long – up to 10cm, thin, charged chain into a small volume. The second
difficulty is how to read the information stored in DNA when it is confined within the
cell nucleus. Thus, in eukaryotic cells, DNA must be compacted sufficiently to fit within
the small confines of the cell nucleus, while the information it contains must be made
available for processes such as transcription, translation, duplication etc. How
compaction and access are achieved in the nucleus in vivo is the focus of much intense
8
research.
The first step in compaction and organization of DNA in the nucleus is the
formation of multiple nucleosomes (Kornberg and Thomas, 1974) along the length of the
DNA. A nucleosome is formed when DNA wraps itself 1.75 times around a core
composed of 8 histones (Luger et al, 1997). The core particle is an octomer of two each
of H2A, H2B, H3 and H4 histones. A fifth histone, H1 or H5, may act to link the DNA
between successive core particles. A schematic representation of the nucleosome is
presented in figure 1.2.
Many different amino acids are present in the tails of the histones but the lysines
are of significance in this work for reasons we will discuss in the next section.
DNA-histone interactions are mainly electrostatic in origin, and result from the
attractive force between the negative charges on DNA and positive charges on histone Nterminal tails. Of the core histones, the helix dipoles in H2B, H3 and H4 form a structure
with a net positive charge in the region of interaction with DNA. All of the core histones
carry a net positive charge in the amino-terminal residue tails (Widom, 1997). Interaction
of DNA with histones resulting in nucleosomes is associated with a stabilization free
energy F ~ 20k BT (Marko and Siggia, 1997).
The next step in the process of folding DNA in the nucleus is formation of
chromatin, which is a higher order structure arranged out of nucleosomes and other
proteins. The structure of the resulting 30nm wide fiber is still being studied (Finch and
Klug, 1976); proposals include a twisted spiral array of stacked nucleosomes, a
disordered zigzag structure (Robinson and Rhodes, 2006), or some other possibility.
9
Histone Core Particles
Nucleosome
DNA
Figure 1.2: A schematic representation of a nucleosome. One can see the eight core
histones as individual blue spheres. The group of eight histones forms the core particle
around which the DNA curves ~1.7 times. The DNA represented by the black curve in
this figure looses ~50nm to when one nucleosome forms. H1 is not shown.
10
1.4 Histone Post-Translational Modifications
Post-translational modifications (PTMs) are alterations made to proteins after
they have been synthesized in the cell. There are many forms of PTMs but the most
common types are acetylation, phosphorylation, methylation, and ubiquitination.
Histones have multiple amino acid side chains that can be modified: arginines are
typically methylated, and lysines are typically acetylated. Acetylation of histones
typically takes place at the lysine residues on histones H3 and H4. Specialized protein
enzymes facilitate this reaction. Histone- acetyltransferases (HATS) help ligate the acetyl
group while histone-de-acetyltransferases (DATS) reverse the process (Petsko et al,
2004). When all or nearly all of the lysines in the N-terminal tail are acetylated, the
histone is said to be hyper-acetylated (Clayton et al, 2006).
Acetylation of histones can lead to modifications of the histone-DNA and interhistone interactions in nucleosomes. Bulk assays have shown that acetylation of
nucleosome core histones reduces the overall stability or binding affinity of the
nucleosome to DNA (Jenuwein et al, 2001, Clapier et al, 2002, Hassan et al, 2001, Tse et
al, 1998, Carruthers et al, 2000). Nevertheless, how or how much acetylation affects the
stability of nucleosome is still not well understood.
Although the stability of nucleosomes may be modulated in a variety of ways,
post-translational modifications – especially in combination with mechanically
destabilizing forces and torques applied by proteins bound to DNA segments near
nucleosomes - may be among the most important mechanisms for changing the binding
11
dynamics of DNA with the octomeric core particle, perhaps leading to local unfolding of
chromatin and thus allowing access to the DNA that was previously occluded.
1.5 Response of a Single DNA Polymer to Micromechanical Manipulation
In physiological conditions - a buffered aqueous salt solution with pH of ~7.5 at
300K - DNA behaves like a three dimensional random walk (Bustamante et al, 1994) on
large enough length scales. DNA in solution is subject to the thermal interactions with the
surrounding medium. The random thermal interactions of solvent molecules produces
kinks and bends all along the length of a free DNA molecule which gives long DNA
polymers a bent and kinked configuration. On short length scales though, shorter than the
~50nm persistence length, DNA behaves like a rigid rod. Thus DNA is referred to as a
semi-flexible polymer: floppy on long length scales, but rigid at short scales.
In physiological conditions, the relevant energy unit is k BT which, at a room
temperature of 300K, is 4.1pN·nm.The configuration of DNA can be modified by applied
forces which if sufficiently large can straighten out the thermal kinks and bends. The
forces required are on the order of ~ 0.1pN (~ 0.1k B T / nm ). In order to affect the
hydrogen and base-stacking interactions holding DNA together, larger forces are needed.
In fact, near ~60pN, the force is large enough to produce a radical restructuring of the
secondary structure of DNA referred to as the overstretching transition.
The first experiments in single molecule biophysics by Smith, Finzi, and
Bustamante focused on measuring the response of a single DNA molecule in
12
physiological buffer to forces in the range of ~0.1pN to ~10pN (Smith et al, 1996). In
later experiments, reversible DNA overstretching where DNA extends up to seventy
percent of its native contour length in response to a force of ~65pN was discovered
(Cluzel et al, 1996, Smith et al, 1996). Other experiments assessed how changing buffer
conditions affected DNA’s mechanical response (Williams et al, 2001).
Figure 1.3 is a force-extension plot for DNA showing its response to various
forces (Cluzel et al, 1996). For forces from 0.1pN -2pN, DNA displays non-linear
entropic elasticity in which thermal kinks and folds along the contour of the molecule are
removed, but the covalent and hydrogen bonds in DNA are not stretched. For forces from
~2pN to ~65pN, linear Hookean elasticity is observed; here DNA actually stretches so
that its contour length changes with force. When the force is ~ 65pN, a reversible
overstretching response is seen. At this force, the molecule’s contour length – the length
of the molecule at full extension - abruptly changes by 1.7 times its zero-force contour
length – for λ-DNA the zero force contour length, i.e. the length of the molecule as
measured along the center line of the double helix, is 16.4μm.
DNA’s mechanical response to an applied force has been studied using various
single molecule techniques, and is very repeatable. In addition, the molecule’s force
response characteristics have been assayed under various salt concentrations, loading
rates, and buffers (Wenner et al, 2002, Williams et al, 2001).
For DNA subjected to forces from ~0.1pN to 20pN, the relation between force
and extension has been shown to follow the theoretical worm-like chain (WLC) model (
Marko and Siggia, 1995):
13
fb
k BT
1
1
4
z
L0
f
K0
2
1
4
z
L0
f
K0
(1).
Here, f is the applied force, k B is Boltzmann’s constant, T is the absolute temperature of
~ 297K, b is the persistence length ~50nm, L 0 is the contour length of 16.4μm, K 0 is the
elastic modulus of DNA given as 1000pN, and z is DNA’s observed end-to-end
extension.
The overstretching transition is not well understood; there exist at least two
competing explanations, each of which has substantial experimental support. One
suggestion has been that the change is contour length is produced by an uncoiling of the
double helix to another quasi-helical structure with a longer helix repeat. Another
explanation revolves around force-induced dissociation (often referred to as DNA
melting) of the complimentary base-pairing holding the two strands of DNA together
leading to the observed sudden extension change.
Another set of experiments explored the mechanical response of DNA that was
twisted and torqued while still subject to tension (Strick et al, 1996). In these
experiments, “plectonomic supercoils” were formed (Strick et al, 1996). With
developments in micromanipulation techniques it has been possible to map out DNA’s
force-extension response to a wide range of twist stresses, torques, and forces. The force
required to pull apart the two strands of DNA’s double helix has also been studied using
single molecule techniques (Essevaz-Roulet et al, 1997). DNA unzipping has been found
to require ~15pN of force.
14
1.6 Response of Single DNA-Protein Complexes to Micromechanical Manipulation
The next development in the field was to study the interaction of DNA with
proteins at the single molecule level. A variety of DNA-protein interactions have been
investigated starting with the role of bacterial lactose-repressor protein in DNA looping
(Finzi and Gelles, 1997). Later experiments studied how RNA-polymerase motored along
DNA in an effort to better understand how DNA is processed within the cell (Wang et al,
1998). These techniques have also been used to study DNA’s interaction with
intercalating agents such as ethidium bromide and proteins such as recA (Vladescu et al,
2005, Leger et al, 1998). Many more results using other proteins have been reported, and
work continues in extending these techniques to study a wider range of DNA-protein
interactions.
One of the earliest works studying the interactions of proteins with DNA was
performed by Leger et al where recA was introduced to DNA under tension (Leger et al,
1998). In this work, they observed the polymerization of recA on the DNA under tension
and the subsequent elongation of the DNA at forces less than 55pN (Leger et al, 1998).
Another early study looked at DNA bending and looping proteins such as HMGB1,
NHP6A, and HU (Skoko et al, 2004).
15
Force (pN)
DNA Overstretching
DNA Hookean Response
DNA Entropic Elasticity
DNA Extension (µm)
Figure 1.3. This force extension plot shows three distinct responses of DNA to applied
tension. First, at low forces less than 2pN, DNA has entropic elasticity. Second, DNA is
subject to Hookean elasticity from ~2pN to ~65pN. Third, one can observe the
overstretching phase of DNA for forces in excess of 65pN. Figure courtesy of
http://alice.berkeley.edu/~steve/DNAstr.html, annotations provided by author.
16
Using vertical magnetic tweezers, the authors found that the amount of
compaction observed and the energy required to re-extend a compacted DNA molecule
was dependent on the protein concentration used. This concentration-dependent effect
was also seen in a later study of the DNA looping protein Fis by Skoko et al (Skoko et al,
2006). They found that when the protein was introduced to the DNA under low forces,
the DNA could completely compact and would only begin to unfold with the application
of ~10pN of force.
RNA polymerases and DNA polymerases process DNA for transcription and
replication, respectively; both have been studied extensively using a variety of molecular
biology techniques (Wang et al, 1998, Shamoo and Steitz, 1999). Using a single
molecule approach, Wang et al found that RNA polymerases were able to move along
and transcribe DNA until the tension on the DNA exceeded a critical force of between
22pN and 25pN. In this study, they also found that the RNA polymerase was capable of
producing 5pN of force along the DNA as it moved along (Wang et al, 1998). Similar
experiments were performed with DNA polymerase by Shamoo and Steitz (1999).
1.7
Response
of
Chromatin,
Nucleosomes,
and
Nucleosomal
Arrays
to
Micromechanical Manipulation
Single molecule micromanipulation studies have also been extended to chromatin
and nucleosomal arrays starting with the pioneering work of Cui and Bustamante, who
presented data showing that chromatin fibers acted like Hookean springs when subject to
17
low forces near~5pN but were irreversibly elongated when subjected to repeated high
force - ~20pN - extensions (Cui and Bustamante, 2000).
Soon after, Bennink et al observed the force-induced dissociation of individual
nucleosomes but found these events at high forces between 20pN and 40pN (2001). Later
work on tension-induced nucleosome dissociations reported more accurate measurements
of the forces under which nucleosomes could remain stable, but were unable to show the
distinct rupture of individual nucleosomes at any force less than 10pN (Brower-Toland et
al 2002, Pope et al 2005). This was important because the expected critical minimum
force f* below which nucleosomes could remain intact under the applied mechanical load
was thought to be in the single pN range.
Another approach was initiated by Yan et al who devised a horizontal magnetic
tweezers system to study chromatin assembly and nucleosome dynamics in the presence
of external tension (Yan et al, 2007). An important advantage of their approach was that
forces could be applied at low loading rates. Their results for f* improved, and they found
f* ≈ 4pN but still higher than the predicted value (Yan et al, 2007). These and other laser
tweezers- and magnetic tweezers-based experiments on chromatin and reconstituted
nucleosomes are described in detail in the discussion section of chapter 4.
The inability to detect nucleosome dissociation for very low forces ~1pN may be
attributed to the limitations of the experimental designs used. In the case of optical traps,
high loading rates may have been the limiting factor. In the case of vertical magnetic
tweezers, the low resolution, optical feedback mechanisms may have prevented detection
of small 50nm extension jumps. With only a few groups using a transverse magnetic
18
tweezer system remotely comparable to the one described here, the main limitation to
observing low force jumps in extension is either the use of an open cell design as in Yan
et al or a single bead design as in Chen et al and Sun et al (Yan et al 2007, Chen et al,
2011, Sun et al, 2008).
1.8 Experimental Methods for Single Molecule Micromanipulation
1.8.1 Optical Tweezers
One technique for manipulating single molecules of DNA is the optical trap, more
frequently called optical tweezers. Laser light is first focused onto ~1 micron diameter
dielectric – usually polystyrene – beads. The momentum transfer that results from the
scattering of photons from the bead results in a well-defined potential well with the bead
trapped at the center.
Although first developed by Arthur Ashkin (Ashkin, 1970) as a tool for trapping
neutral atoms, optical tweezers have become the primary tool used by single-molecule
biophysicists since the pioneering works of Carlos Bustamante in 1992 and Steven Block
in 1993 (Svoboda et al, 1993, Smith et al, 1992).
DNA is prepared with a bead at one end while the other end is chemically fixed to
a glass surface. The polystyrene bead can be seen with an optical microscope and trapped
in the laser spot. By holding the glass surface fixed while moving the laser spot, the
DNA’s extension can be varied as one pleases. The resulting force can be measured in a
variety of ways, usually by detecting how the Brownian fluctuations of the trapped bead
19
change with changing force. Variations on this arrangement include using beads at both
DNA ends and then using dual traps to manipulate both beads, and a combination of a
single trap for one bead and micropipette aspiration for the other.
This technique initially gained favor with single molecule experimentalists
because of its ability to precisely manipulate the small dielectric beads with minimal drift
and apply a wide range of forces between ~1 to ~ 100pN to DNA. It has the ability to
precisely control the position of the trapped particle allowing “fixed-extension”
micromanipulation experiments. It also has the ability to manipulate the dielectric bead in
the plane of focus allowing simultaneous visualization of the trapped beads when two
beads are used.
This approach also has some limitations. The laser light used to trap dielectric
beads can destroy the biological sample, and DNA is especially susceptible to radiative
damage. The electro-mechanical complexity of these systems makes them extremely
difficult to set up and complicated to use, and makes it highly sensitive to mechanical
noise. If control over the applied force is desired, then substantial modifications to the
system have to be made that bring their own complexities and limitations.
In figure 1.4, a dual laser beam optical trap is depicted. Other variations of the
optical trap exist, but this highlights the most complex and the most versatile form of
laser tweezers available.
20
Dielectric Beads
DNA
Focused Laser Light
Figure 1.4: A schematic of a dual laser optical trap. This shows the principle behind the
operation of an optical tweezer system. The dielectric beads are held in place by a
potential well created by the focused laser light. The DNA is tethered on both sides to
dielectric beads. By either adjusting the focus of the laser beams or by moving the laser
beams,
DNA’s
end-to-end
extension
can
be
adjusted.
21
1.8.2 Atomic Force Microscopy (AFM)
Atomic-Force-Microscopy,
AFM,
is
another
popular,
single-molecule
micromanipulation technique. AFM was initially developed by Gerd Binnig and Heinrich
Rohrer in 1986 as an alternative to scanning tunneling microscopes (STM) (Binning et al,
1986). This technique utilizes a cantilever with a sharp tip and a piezo-electric stage. A
schematic of this system is shown in figure 6. One end of a biopolymer is attached to the
probe on the cantilever; the other end is attached to the moving piezo-electric stage. As
the stage is slowly moved, the biopolymer translates this movement to the cantilever. A
laser that reflects off the cantilever provides a means of measuring the force acting on the
taught polymer. Attributes include the ability to apply forces from 10 to 10000pN, the
ability to work in physiological buffers, and the ability to resolve sub-nanometer spatial
deflections. It is especially useful when studying the strength of individual molecular
bonds and the folding energies of proteins.
The major limitations of AFM come out of the complex mechanical nature of the
device. The cantilever, although micron in scale, is large in comparison to the molecules
it observes. This large cantilever is inherently stiff and is incapable of applying low level,
0.01 to 10pN, forces. Also, because two ends of a biopolymer need to be precisely
attached to both the movable stage and the force probe, it is difficult to perform many
experiments in a reasonable amount of time.
22
1.8.3 Vertical Magnetic Tweezers
Magnetic tweezers take advantage of the force that a superparamagnetic particle
feels when placed in a changing magnetic field. In fact, the force is known to be
proportional to the spatial rate of change of the field, so that the regions with magnetic
field lines spaced closely together produce a larger force than regions where the field
lines are spaced farther apart. In these systems, one or more magnets are arranged so that
the resulting field changes rapidly in one direction – which will be the direction of the
applied force – but is constant in the other two directions. This ensures that the force
points along a single line, without off-axis components.
Magnetic tweezers have many advantages which we will discuss in more detail
below (together with some drawbacks.) In general though, they are, in most cases,
mechanically simple and require only the interaction of a magnetic particle with a
changing magnetic field.
In most magnetic tweezers (MT) the system is constructed in a vertical
arrangement starting with a magnet or system of magnets, descending down to a flow cell
containing the sample, and finally reaching a high-power microscope objective. Inside
the flow cell, one end of a DNA molecule is fixed to a biochemically prepared cover slip
while a superparamagnetic bead is attached to the other end. The magnet positioned
above the DNA construct applies force on the bead through the interaction of the bead
with the changing magnetic field. The magnetic field gradient pulls along a vertical axis,
pulling the super paramagnetic bead vertically off of the cover slip. The bead’s vertical
23
movement is monitored by a high-power microscope objective. With changing extension
of the DNA tether, the bead undergoes displacement from the focal plane of the
objective; by refocusing the objective by traversing it through a known distance, the
change in tether extension can be determined. A schematic of a typical vertical magnetic
tweezer is given in figure 1.5.
The idea of using magnets to study DNA first arose in the lab of the Crick and
Hughes in 1950 (Crick et al, 1950), but it was the work of Smith et al in studying the
effect of DNA binding drugs on the elasticity of DNA that cemented its status as a
formidable approach to the micromanipulation of single DNA molecules (Bustamante et
al 1994). Later, in an important development of this approach, Strick et al attached the
DNA to a cover-slip and a magnetic bead in such a way as to limit its ability to freely
rotate (Stick et al, 1996). By ligating both 3’ and 5’ termini of the DNA to a bead at one
end, and similarly the 3’ and 5’ termini at the other end to a glass surface, they made it
possible to apply torque to the DNA molecule. By rotating the magnets above the sample
cell, the DNA could be simultaneously twisted and have a force applied to it. With this
setup, Strick et al could form supercoils along the length of the taut DNA molecule and
study their effect on DNA elasticity.
Vertical magnetic tweezers are limited by the need to continually refocus on a
bead that moves in and out of the focal plane. The optical-mechanical feedback loops
required severely limit the spatial and temporal resolution of such setups. If many
changes in DNA’s extension occur during an experiment, the feedback mechanisms can
neither react quickly enough nor with enough precision to make nm scale measurements.
24
N
S
S
N
R
F
Force
pulling
upward
Figure 1.5: A schematic of a vertical magnetic tweezer. From the top, one can see the two
magnets aligned in a north-south, south-north configuration. This arrangement provides
an exponentially changing magnetic field gradient close to the magnet. Next, one can see
the DNA tethered to both a magnetic bead and the cover glass. The magnetic field
gradient acts to pull the beads off the floor. Below sits a high power objective to actively
track the changes in the end-to-end extension (z).
25
1.8.4 Transverse or Horizontal Magnetic Tweezers
Horizontal magnetic tweezers have been developed by Yan et al (2004) to
overcome some of the deficiencies in existing designs of vertical magnetic tweezers, but
their design has some limitations too. In their approach in-plane micromanipulation of a
DNA-tethered bead pair consisting of a magnetic and non-magnetic bead was achieved
using an open cell design in which an aspiration micropipette entering at an angle was
used to capture the polystyrene bead while a small magnet applied tension on the
superparamagnetic particle (Yan et al, 2004). Chen et al introduced a modification of the
method used in vertical tweezers by coating the inside wall of a square capillary tube that
would then be moved in relation to a magnet in the same plane (Chen et al, 2011). Taken
together, these works mark an important development in magnetic tweezers design, and
significantly enhance their utility for biophysical experimentation, though they also
suggest areas of improvement. For instance, in Yan et al’s system, the experiment takes
place in a cell that is open to the environment. This allows noise from the environment to
affect the experiment. Also, because of the open design, buffer evaporation, though
mitigated, is a major problem. Chen et al’s design is still subject to limited amounts of
force. Others have used transverse sample cells but have been limited in the amount of
available force and freedom of movement (Sun et al, 2008).
26
1.9 Research Problem and Approach
Histones can undergo many types of post-translational modifications (PTMs);
one, described earlier, is acetylation. Being critical to the storage of genetic information,
numerous labs have tried a variety of different approaches to ascertain what effect histone
acetylation has on the stability of nucleosomes. In particular, some experimental support
has been obtained for the hypothesis that nucleosomes constituted out of PTM-histones
are less stable and more prone to (partially) release the bound DNA.
This mechanism could be further facilitated by the presence of mechanical tension
on the nucleosome. Many protein machines, such as DNA and RNA polymerases, bind
DNA and act processively (sequential movement along the DNA) on it in regions
populated with nucleosomes while applying large forces in the ~0.1pN to ~10pN range.
The possibility of PTM-induced nucleosome instability and the presence of forceapplying protein machines in the cell nucleus leads to two questions that we address in
this work: 1) How do post-translational modifications of histones affect the stability of
the nucleosomes? and 2) How do nucleosomes formed with native and posttranslationally modified histones respond to applied tension?
Before addressing these two questions, we need to first establish the force
required to destabilize a nucleosome. In the absence of external tension, nucleosomes will
form spontaneously when all components are present. In fact, this process is driven by a
fairly large free energy gain
F ~ 20k B T (Marko and Siggia, 1997) resulting mainly
from the strong electrostatic interactions that stabilize the DNA-histone complexes into
nucleosomes.
27
What force f will induce a nucleosome to dissociate? This can be calculated by
noting that if the nucleosome dissociates, the work done is f l where l is the length of
DNA previously coiled up, about ~50nm. If this work is larger than
F, then
nucleosomes will be destabilized. Thus, when the applied force is such that f* l =
F,
i.e. when the external work done in increasing the DNA extension by l is equal to the
free energy gain from forming a nucleosome, nucleosomes will become marginally
unstable, with equal proportions of bound and dissociated nucleosomes in thermal
equilibrium. From the values give for l and F, it follows that f* ≈ 1.6pN.
To verify this hypothesis, we have developed a single molecule assay to
determine f* that overcomes many of the limitations of previous studies that have tried to
verify f*, but have instead found it to be much larger than what was predicted. Our
approach also allows us to characterize how f* is affected for native nucleosomes and
those constituted out of acetylated histones.
We then consider how acetylation affects the response of nucleosomes to force.
Our single molecule technique based on a novel transverse magnetic tweezers design
allows us to observe individual nucleosome pop-offs for forces > f* by tracking the
minute ~50nm changes in DNA extension that result when an individual nucleosome
dissociates. Based on these data, we characterize in detail the distributions in step sizes,
so that we can tell what proportions correspond to single nucleosome unbinding – ~50nm
steps-, two nucleosome unbinding ~100nm steps -, three nucleosomes unbinding ~150nm steps -, and so on, and to compare the results for native and PTM-modified
nucleosomes. We also study the distribution in f* that characterizes nucleosomes, finding
28
that some nucleosomes have much larger f than the majority of the population, and
*
establish how these distributions change upon histone acetylation – which they do
profoundly, as discussed in detail in chapter 4; also see below for a very brief summary.
The distribution of F is also mapped by directly calculating the work done is
dissociating nucleosomes from our force and extension data for both kinds of
nucleosomes. Going beyond equilibrium quantities like f* and F, we have also been able
to determine the time required for individual nucleosomes to rupture under applied
tension, and to establish the distribution in those times for both kinds of nucleosomes.
As far as we know, this is the first time any one has confirmed that f* ≈ 1.6pN,
and reported data for distributions of step lengths, critical forces, free energies, or kinetic
time constants for both kinds of nucleosomes under such a wide range of forces,
especially for forces between 1 – 10pN. It is important to emphasize that by the very
nature of these quantities, it is difficult to see how alternative techniques based on bulk or
ensemble measurements could be devised to directly obtain these kinds of data.
The critical requirement for obtaining these data was to develop a low noise
single molecule force transduction system capable of applying a wide range of forces
quasi-statically. Previous attempts to study nucleosomes could not establish f* accurately
or obtain the distribution in step sizes, critical force, or binding affinities because they
suffered from either high force loading rates or high mechanical noise which prevented
detection of steps, especially those at forces ~ 1pN. We report on the development of a
horizontal magnetic tweezers that solves these problems and allows us to obtain for the
first time high spatial resolution nucleosome pop-off step data at very low forces (even
29
for forces < f ). Using a semi-closed sample cell to reduce mechanical noise, a 40X
*
microscope objective and sophisticated image processing techniques to resolve individual
steps, our approach takes the best aspects of all other techniques used to date while
minimizing the limitations associated with them.
1.10 Results
In our setup, we use a sample chamber that is sealed on five of its six sides, the
open side allowing an aspiration and protein microspray pipette to enter horizontally.
Within the chamber, a single bar magnet produces a magnetic field gradient acting in the
focal plane of an objective. We use a two-bead DNA construct that allows precise
measurements of DNA’s end-to-end extension. Using aspiration on a polystyrene bead
attached at one end of the DNA, we can hold the DNA construct above the objective.
Meanwhile we can very slowly and precisely adjust the position of a simple bar magnet
to pull on a super-paramagnetic particle attached at the other end of the DNA molecule,
thereby applying tension over a wide range of forces while monitoring the DNA’s end-toend extension. As a result, we achieve force loading rates of 0.008pN/s, perhaps three
orders of magnitude better than what has been previously reported.
After extensive validation of our micromanipulation system on single DNA
molecules – see chapter 2 -, we study tension-modulated stability of native and posttranslationally-modified nucleosomes reconstituted on -DNA. In chapters 3 and 4, we
discuss our observation of the predicted theoretical critical force of 1.6pN first proposed
30
by Marko and Siggia in 1997. We experimentally determined the distribution of jumps in
extension – steps – associated with single or multiple nucleosome disruptions and find
that for both native and PTM histones the distribution has a prominent peak at ~50nm
with a clear secondary peak at ~100nm emerging for PTM nucleosomes. We find that as
a function of force the distribution of steps of any size – 50nm, 100nm, etc – is peaked at
a low force of ~ 1.6pN for both native and PTM nucleosomes, but show that, in
comparison to PTM nucleosomes, the distribution for native nucleosomes is much
broader with many steps – of any given size – occurring at forces from 1.6pN up to
20pN. We experimentally determined the differences in the binding free energy of native
and PTM nucleosomes and show an ~ 11k B T offset in both the average and median free
energies over a range of forces from ~1pN – 25pN, providing for the first time a single
molecule measurement of the reduction in nucleosome stabilization free energy upon
histone (hyper) acetylation.
As discussed in detail in chapters 3, we have observed, for native nucleosomes, a
distinct pattern of response to forces in three ranges: less than 2.5pN, 2.5pN - 12.5pN,
and greater than 12.5pN. For forces less than the critical force, we show that nucleosomes
are in equilibrium; we find that they rupture at a steady rate up to a force of ~ 12.5pN
followed by rapid dissociation for forces greater than 12.5pN. Moving on to inter-step
time interval measurements (which are related to the kinetics of nucleosome pop-offs),
we present data showing the lack of a difference in the average time interval between
successive nucleosome dissociation events for f > f* is the same for both native and PTM
31
nucleosomes, with average time between events of ~5s. We show that the median rates in
rupture events are also similar – both ~ 2.5s - but much less than the average of ~ 5s.
1.11 Plan of the work
This dissertation is structured as follows. In chapter 2 our horizontal magnetic
tweezer design is discussed, including detailed descriptions of DNA force-extension and
calibration experiments. In chapter 3, DNA-histone binding experiments using native
histones are discussed. In chapter 4, DNA-histone binding experiments using PTMhistones are discussed and a comparison is made between reconstituted native and PTM
nucleosomes. In chapter 5, a summary of the previous chapters is provided followed by
discussion of some possible future directions for experimental work.
Chapter 2
Horizontal Magnetic Tweezers
In this chapter, I describe the transverse magnetic tweezers system I have
developed for carrying out single molecule experiments on DNA and nucleosomal arrays.
I was guided by the requirement for a piconewton-scale force transduction system that
could apply forces quasi-statically, and with the ability to monitor changes in DNA
extension with very high resolution to allow detection of single nucleosome unbinding
events. I begin by describing in detail here and in the appendix the design of our
tweezers, the biochemical protocols, and the associated data processing steps, and then in
the following sections discuss results from various calibration tests I have performed to
establish and validate the operating characteristics of our tweezers. Portions of this
chapter have been submitted in manuscript form and are under consideration for
publication in a research journal.
2.1 Description of System
Figure 2.1 shows the basic principle of our transverse magnetic tweezer. Linear λDNA is prepared with a non-magnetic, polystyrene bead on one end and a
superparamagnetic bead on the other. The polystyrene bead is held in place by an
aspiration pipette with a 2-2.5µm opening. A force, in the range of 0.05pN -100pN, can
be applied to the superparamagnetic bead by adjusting the distance between the magnetic
32
33
bead and a small bar magnet. By design, the rate of change in the magnetic field as a
function of distance from the magnet, henceforth referred to as the magnetic field
gradient, and thus the force, is in the focal plane of the objective midway between the
floor of the cell and the top of the magnet.
Experiments are performed in a semi-closed chamber made using two #1 coverslips (Fischer Scientific), cut glass slides, and a 4mm x 2mm x 1mm neodymium bar
magnet (Indigo Instruments). See figure 2.2. The cover-slips form the floor and ceiling,
while the cut sections of regular microscope slides form three of the four walls of the
chamber. The cut glass and the bar magnet are both glued to the bottom cover-slip with
clear RTV silicone sealant. The fourth side of the chamber is purposefully left as an
opening. This opening, a 1mm x 40mm rectangular breach, is designed to allow
horizontal insertion and movement of aspiration and other micropipettes.
The chamber is mounted on an aluminum stage. The stage has an appropriately
sized opening that allows the objective to come arbitrarily close to the underside of the
floor of the chamber, and is attached to an Eppendorf 5171 motorized three axis
micromanipulator with a 0.160µm step size and a minimum step rate of 0.320µm/s. The
chamber is secured to the stage using clear, double- sided tape (3M). In order to make
fine adjustment of the aspiration pipette over the objective, it is clamped into the arm of a
hydraulic, three-axis micromanipulator (Siskiyou). All micromanipulators are built
around a Nikon Dyaphot inverted, light microscope. The Eppendorf 5171 is mounted on
the microscope’s side bench and positioned to allow easy movement of the stage above
the objective. The hydraulic manipulator is mounted on a carriage designed to bridge a
34
Aspiration Pipette
m
F
y
x
N
S
z
Not to scale
Figure 2.1: A glass micropipette (~1.5µm to 2.5µm opening) and a bar magnet (1mm x
2mm x 4mm) form the basis of our magnetic tweezer. An origin is defined at the center
of the inward facing 4mm x 1mm magnet wall. The long axis (4mm) of the magnet is
positioned parallel to the aspiration micropipette. The bead-DNA-bead constructs are
formed by attaching a non-magnetic bead (3.0µm) to one end of a λ-DNA molecule and a
superparamagnetic bead (2.8µm) to the other. The non-magnetic polystyrene bead is held
by aspiration while the superparamagnetic particle is placed in the magnetic field with the
gradient of the field delivering the force on the magnetic bead. By adjusting the distance
(not to scale) to the defined origin, forces ranging from 0.1pN to 100pN can be realized.
In this figure, the plane of the paper is the plane of focus.
35
rail system (Grainger) that allows it to easily move horizontally into the chamber and out
of the way when not needed or when loading the pipettes.
The stage is positioned so that the semi-closed chamber is above a 40X, 0.65NA,
bright-field objective (Leitz) with a working distance of 500μm. Labview allows us to
stream and record videos captured by a CCD camera (Sony XCD-U100). An additional
zoom lens (Edmund Scientific) lies in the optical path between the objective and the
CCD. I record 8-bit grayscale images at 15 frames per second. The data from experiments
are stored as movies on an external hard drive (CFI), and analyzed on a Windows PC (HP
Compaq DX 7500 Micro Tower). DNA extensions and forces are calculated offline using
our own particle tracking and localization software written in MATLAB.
2.2 Preparation of DNA, Beads, Buffer Chamber, Pipettes, Buffers, Protein
Solutions
A non-magnetic, 3.0µm diameter, polystyrene bead, (Polysciences) is used for
aspiration, while force is exerted on a superparamagnetic, 2.8µm diameter microsphere
(M-280, Dynal Biotech). The bead has saturation magnetization of 10 Am2/kg (mass) and
14 kA/m (volume) and remains saturated at the distances from the magnet used for our
experiments. Polystyrene beads are labeled with anti-digoxigenin using standard
protocols (Skoko, 2006). Magnetic beads come pre-coated with streptavidin.
36
m
y
x
z
Not to scale
Figure 2.2: Experiments are performed in a semi-closed chamber custom built using two
#1 cover-slips, cut microscope slides and a 1mm x 2mm x 4mm bar magnet. The coverslips form the floor and ceiling of the semi-closed chamber. The cut pieces of microscope
slides form three chamber walls. The glass pieces and the magnet are fixed to the bottom
cover-slip. From the side left open, I can introduce the aspiration pipette to capture beadDNA pairs. Independent 3-axis micromanipulators (not shown) are used to position the
pipette and the cell above the objective. The pipette and the sample cell move
independently over the 40X, 0.65NA microscope objective. Figure not to scale.
37
I use linear, ~50kbp, λ-DNA molecule (New England Biolabs) with 15 base
overhangs on each end: 3’-end overhang is gggcggcgaccg, and 5’-end overhang is
aggtcgccgccc (Smith et al 2006). These ends are complementary to single-stranded DNA
oligomers with conjugated biotin (3’) and digoxigenin (5’) chemical tags (Skoko, 2006,
Haber and Wirtz, 2000). The oligomers are ligated to λ-DNA using T4 ligase following
standard protocols (Skoko, 2006). The resulting DNA molecules can thus bind
streptavidin coated, superparamagnetic beads on the 3’ end, and antidigoxigenin-coated
beads on the 5’ end.
DNA-tethered beads are prepared by incubating labeled superparamagnetic and
polystyrene beads with end-functionalized λ-DNA. Incubation is carried out in 1X TrisEDTA solution at 28ºC to 30ºC overnight in an incubator (Bambino). Incubation takes at
least 8 hours (Yan et al, 2004), but I have found that 14 hours yields more tethered
DNAs. These steps a described in detail in the appendices.
The micropipettes needed for aspiration are formed in a two step process. First,
standard 1mm, thick-walled, glass capillaries (World Precision Instruments) are pulled in
a horizontal micropipette puller (Sutter Instruments model P-97) leaving submicron
barbs. These pulled barbs are then cracked and made useable using a microforge
(Sensaur, de Fonbrune style microforge). The resulting cracked and polished
micropipettes are left with an opening of approximately 1-2.5μm estimated based on a
comparison to the aspirated polystyrene bead. These steps are described in detail in the
appendices.
38
The pipette is filled with buffer prior to being connected to a syringe and syringe
pump (New Era Pump Systems model NE-1000). This completely-fluid-filled system can
then easily and consistently aspirate polystyrene beads. When a bead has to be aspirated,
the syringe pump is set to a withdrawal rate of 5.5mL/hr – 7.5mL/hr. This creates a back
pressure at the mouth of the pipette which holds the bead in place.
For details about buffer preparation and protein purification steps, see appendices.
2.3 Data Analysis
2.3.1 DNA Tether Extension Measurement
All of our DNA extension data is analyzed with a bead tracker algorithm that
identifies the centroids of the tethered beads. MATLAB code identifies the beads and
traces their movement during an experiment. The bead tracking software tracks the
centroids of the Airy disks of the two beads and allows calculation of their separation in a
straightforward way. The Airy disks are the pattern produced by the light diffracted by
the bead. The positions of the beads are recorded as a function of time. Their separation is
calculated as the vector distance from the aspirated bead to the magnetic bead. The
algorithm is designed to detect the two beads in each frame based on the following
criteria: (1) their diameters, the polystyrene bead is 3.0µm and the superparamagnetic
bead is 2.8µm and (2) their position in the field of view in relation to the origin in the
upper left hand corner of the frame. In all of our experiments, this would lead to the
39
polystyrene bead being detected first, when analyzed from the top left to bottom right,
and the superparamagnetic bead second.
With the current computational techniques, the system can resolve changes in
extension far beyond the optical limit – currently achieving 10 nm extension changes.
The current system identifies the diffraction pattern and the gradient of the diffraction
pattern of each bead and identifies the edges of these patterns. After MATLAB identifies
the edges of each bead, it computes the centers of each. As the edges move, so do the
centers. MATLAB then tracks the movement of the each of these centers. By monitoring
the separation of the centers, the system can resolve minute changes in the extension of
the DNA molecule.
2.3.2 Force Measurement by Fluctuation-Dissipation Theorem
Thermal fluctuations of the centroid of the magnetic bead transverse to the
direction of force are used to calculate the force acting on the DNA by using the
following fluctuation dissipation relation (Yan et al, 2004) between the force f z and
average extension <z>:
fz
k BT ( z
/
( x) 2 )
(1)
<(δx)2> is the transverse fluctuation of the magnetic bead, k BT
derivation of equation (1) can be found in the appendix.
2.3.3 Measurement of Bead Velocities
4.1 pN nm . A
40
A modified bead tracker algorithm was used to identify and track beads used in
the calibration experiments. This algorithm identified and counted beads upon release
from the pipette tip. Upon identification, a bead’s position was determined and its
movement from frame to frame was tracked. The magnet, being fixed in position, created
a unidirectional line of force; therefore, this means the beads should only move in one
direction. As the beads moved from frame to frame, the change in distance would be
recorded. Upon reaching the bottom edge of the screen, just before leaving the frame, the
last position would also be recorded. A terminal velocity would be calculated at this point
from 1) the distance traveled from the pipette tip to the bottom edge of the frame and 2)
the time as a function of frame rate. This terminal velocity is then put in the Stokes drag
force equation to compute the force at set distances from the magnet. This technique
assumes that the magnetic field changes little over the distance recorded in each frame.
Therefore, the resulting force acting on a superparamagnetic particle, over the ~50µm
observable in each frame does not change appreciably. This is true at distances of ~
400μm to 500μm and more from the magnet.
2.3.4 Detection of Steps
Using a novel step-finder algorithm, the nucleosomal rupture events were
identified in the extension time-trace data. MATLAB code was developed to identify
jumps in extension within the time traces. Each individual data set is filtered using a
41
bilateral Gaussian kernel-type filter. Then, extensions are binned to form a distribution of
extensions with bin height proportional to the number of successive frames that an
extension occurs in the filtered data set. The second derivative of this distribution is
computed at each bin location, and the extension histogram is added to it. The result is
then squared to avoid negative values; the local minima of the resulting distribution
correspond to steps or jumps; the peaks correspond to the plateaus or constant extension
regions between successive jumps.
The algorithm has been validated in three ways. After applying the step-finder
routine to a data set, we check “by hand” that the steps identified by the routine indeed
correspond to jumps in the data. While this assures us that we are not counting steps that
are not there, in order to determine the false negative rate – how many steps the routine
misses - , we have computationally generated “synthetic” data sets where a fixed number
of steps are placed at known locations in otherwise noisy monotonically-increasing
functions of “time” - which simulate our extension-time traces -, and then applied the
algorithm to it. Since we know how many steps there are as well as other step parameters,
we can check to make sure that the step finder does not identify steps not there and does
not miss existing steps. In addition, since these “steps” are generated artificially, we can
iterate this procedure many times while altering the step size and spacing to gain
confidence in our fitting procedure. We also cross check the results for various
calculation based on the step finder algorithm with known experimental data where
available as another method of calibration. Our algorithm has performed very well
against these types of tests.
42
2.4 Calibration
2.4.1 Extension Calibrations – Graticule
I used both a simple optical graticule and a NIST traceable optical grid – a grid on
a microscope slide with lengths verified at NIST - to define an observed grid length to a
known pixel length ratio to be used in the MATLAB algorithms to calculate forces and
extensions of DNA. This ratio was determined by taking a snapshot of the optical grid
pattern with our Labview frame grabber software and counting the pixels in Photoshop
corresponding to specific distances on the grid. For example, I could count 50 pixels
(each 1μm X 1μm) in an image of a 40μm line in the grid and calculate a ratio of
40μm/50μm to get a ratio of 4/5. This procedure was done before I began my DNA forceextension and protein-binding experiments. This was not done each day, but only when
any changes were made in the optics of the system, i.e. the objective was changed, the
zoom was adjusted etc. With this calibration, I can claim that I am observing single
molecules of DNA under tension because I could then plot the calculated DNA force
extension curve against an empirically verified worm-like-chain model.
2.4.2 Force Calibration - Stokes Law Measurement
Force calibration was performed as follows: using the same chamber described
earlier and a micropipette with a 15μm – 20μm opening, I released free, un-tethered,
43
magnetic beads into either a CaCl2 or glycerin solution. The buffer solutions used, a low
viscosity, 1.5centi-Poise (cP), 25% w/v CaCl2 solution and a high viscosity, 7cP 55% w/v
glycerin solution provided neutral buoyancy for the magnetic beads and much more drag
than standard 1X Tris-EDTA buffer. Viscosities were measured using standard
viscometric techniques.
The results of the calibration experiments are shown in figure 2.3. Typical
calibration experiments covered distances from 300μm to 2500μm from the magnet,
which corresponded to forces of ~100pN to ~0.1pN respectively. I released 30-70
magnetic beads in 100μm increment steps from the magnet and recorded their movement
toward the magnet. They immediately began to accelerate and reached terminal velocity
as they exited the screen ~50μm from the pipette opening, ~ 1 to 2 seconds after release
depending on release position.
The magnetic field gradient would not vary too greatly over the distance of ~
50μm, the z- field of view; therefore, I used the measured terminal velocity in Stokes
drag law (Brodkey, 1967) to evaluate the force at that location:
fz =
6 vz d
(2) .
Here η is the viscosity of the medium, vz is the velocity in the direction of the force, and d
is the bead diameter: d = 2.8µm. Because the velocity of the beads was on the order of
10μm/s, use of Stokes force law was valid because these experiments were all performed
in the low Reynolds number regime. Likewise, the effect of the two vertical bounding
surfaces was negligible because particle trajectories were confined to a plane 500μm
from either.
44
Figure 2.3: In this figure I compare the forces measured using the fluctuation dissipation
theorem with the forces obtained from our calibration experiments using Stokes law.
Magnetic beads were introduced at various distances from the magnet and allowed to
reach terminal velocity, which was measured using video data analysis. Stokes law was
used to calculate the drag force on the bead which allowed us to measure the magnetic
force on the microsphere as a function of distance from the magnet. The thin solid line
shows the average of 6 force-extension experiments. The other lines in the figure
represent calibration experiments in which beads were released into either calcium
chloride or glycerin. The calibration and force-extension measurements agree very well
up to ~35pN, but calibration measurements are unreliable beyond these forces because
magnetic microspheres do not reach terminal velocity within the field of view of the
objective. I see the force increase as ~1/z4; as expected for a magnetic dipole.
45
2.5 Fit to WLC with correct persistence length
Figure 4 presents the results of typical force-extension experiments plotted against
the modified wormlike chain model of DNA (Marko and Siggia, 1997):
fb
k BT
1
1
4
z
L0
f
K0
2
1
4
z
L0
f
K0
(3).
Here, f is the applied force, k B is Boltzmann’s constant, T is the absolute temperature of
~ 297K, b is the persistence length ~50nm, L 0 is the contour length of 16.4μm, K 0 is the
elastic modulus of DNA given as 1000pN, and z is DNA’s observed end-to-end
extension. All experiments are performed in 1X (1mg/mL) Tris-EDTA buffer at room
temperature (~25°C or 297K). All experiments begin when I start recording a movie. I
only start recording after the tethered pair is in position along the perpendicular bisector
of the magnet. Once in position, I note it as the starting point of the experiment. I then
either begin moving the magnet toward the construct or away from it at a rate of either
0.320μm/s or 1.6μm/s. 0. 320μm/s (or 2steps/s) is the smallest step rate that the
micromanipulator can achieve corresponding to two 0.160μm steps each second.
Alternatively, the distance between the magnet and magnetic particle can be adjusted at a
rate of 1.6μm/s (or 10steps/s) and still maintain near equilibrium conditions (the flat
portion of the calibration <20pN curve of figure 2.3). The faster rate is sometimes useful
since this can reduce the overall duration of experiments.
In Fig. 2.4 I show that I can recapitulate DNA’s mechanical response covering
both the entropic - ~0.1 to ~1pN - and elastic – ~1 to ~10pN - regimes. In the main plot,
46
the data were obtained by moving through a force range of ~0.05pN to ~ 20pN at a rate
of 1.6μm/s over the course of 45 minutes, which clearly demonstrates that I can resolve
forces as low as ~0.05pN. In addition, because I cover a very large force range
(equivalently tethered pairs are brought from ~2500μm from the origin to ~500μm
corresponding to the area plotted in figure 3) nearly quasi-statically, individual forceextension experiments can last several hours. I find that the bead aspiration and other
experimental conditions like buffer volume inside the chamber can be stably maintained
for this duration.
In the inset in figure 2.4 I provide data from three experiments focusing on the
force range where DNA’s mechanical response transitions smoothly from entropydominated response to the Hookean response. These data were obtained from
experiments performed at step rates of both 0.320μm/s and 1.6μm/s. The stars and
squares are from experiments performed at 0.320μm/s while the diamonds represent an
experiment performed at 1.6μm/s. No observable dependence of the data on step rates
over this range can be seen, so that equilibrium is maintained over these loading rates. I
also see excellent agreement between the force-extension data and the worm-like-chain
model of polymeric DNA across the range of forces for which the model is valid.
2.6 Typical Low-Force-Extension Experiment for DNA
A typical force-extension experiment is performed as follows: first, I position the
chamber over the objective and then locate the center of the magnet face that is pointing
47
Figure 2.4: Results from several separate DNA force-extension experiments performed
using λ-DNA in our horizontal magnetic tweezer. By positioning our magnet far from the
bead-DNA constructs (> 2000µm), I can apply low forces of ~0.5pN and work within
DNA’s entropic response regime. By bringing the magnet closer (<1200µm) to the
captured pair, I are able to realize higher forces (~5pN to 50pN) and experiment in
DNA’s elastic regime. The solid line represents the wormlike chain theoretical model for
DNA’s response to force. The diamonds in the main plot represent one force-extension
experiment moving from a low force of ~0.5pN to ~20pN at a rate of 1.6µm/s. The inset
shows a zoom-in of three separate force extension experiments at the transition from
DNA’s entropic response to DNA’s elastic response. This shows our ability to maintain a
near equilibrium force transition from entropic to elastic response.
48
into the volume of the cell as shown in figure 2.2. It is important to locate this point since
it is along a line passing through this point that the external force is simultaneously
perpendicular to the magnet and in the x-z plane. Next, the aspiration micropipette is
positioned in the field of view parallel to the long axis of the magnet with its opening
along the line through the origin. Often, during the course of an experiment, I check for
mechanical drift by stepping back to the face of the magnet to see if any unwanted
movement has occurred. With this arrangement, I have a distance-dependent, fixed-force,
magnetic force transducer.
Tris-EDTA buffer and the bead-DNA constructs are then introduced. The latter
settle to the bottom of the chamber very quickly. The second cover-slip is then placed on
the prebuilt and positioned glass cell resulting in a buffer-filled 500μL - 800μL, 25mm x
20mm x 1mm, five sided chamber. Surface tension from the buffer is more than adequate
to form the remaining wall while also allowing micropipettes to penetrate and enter.
Buffer evaporation from this exposed side is negligible under typical sample illumination
conditions, and over a time scale of hours.
I then look for DNA-bead complexes on the floor of the chamber. I find that the
magnetic particle in a pair consistently orients itself closer to the magnet than the
polystyrene bead, giving us a way of distinguishing between the two otherwise visually
identical beads. After identifying a bead pair, I move the micropipette tip behind the
polystyrene bead and begin aspirating. The construct is then brought into position along
the force line described above. The applied force can be adjusted from 0.1pN to well over
50pN of force by changing the distance between the superparamagnetic particle and the
49
magnet (greater force is possible but difficult to utilize with the current system). The
captured pair is known to be positioned along the force line when the bead-DNA
construct is perpendicular to the aspiration pipette and when both beads are
simultaneously in the focal plane.
In figure 2.4, I have shown that DNA can be modeled as a worm-like chain with a
persistence length of ~50nm. The contour length of our DNA is 16.4µm and figure 2.4
also shows the fit to the worm-like chain with this length in equation 3 above.
DNA in physiological conditions under repeated equilibrium changes in force
should exhibit little to no hysteresis. In Figure 2.5, I see the results of an experiment
designed to show the minimal amount of hysteresis in DNA’s response to slow increases
and decreases in the force. As the data show, there exists minimal hysteresis along the
force extension curves produced by stepping at a rate of 0.320μm/s from a low force of
3pN through 50pN and back again. This figure shows the ability of our tweezer to
maintain equilibrium conditions throughout the entire experiment, which lasted ~ 1.5
hours, moving at 0.320μm/s. This quasi-static environment was produced using a loading
rate of between ~0.008pN/s and ~0.1pN/s. The loading rate varies slightly over the
course of an experiment because, as the tethered pair comes closer to the magnet, the
distance dependent force increases much more rapidly as one gets nearer - <1000µm - to
the magnet. At distances greater than 1200µm, a set rate of approach yields an
approximately constant loading rate because the force follows the essentially flat portion
of the ~ 1/z4 curve. I have verified that there is minimal hysteresis when the experiment is
performed at 1.6µm/s (data not shown).
50
This behavior shown in figures 2.4 and 2.5 recapitulates the known response of
DNA to applied force and proves that the novel, transverse magnetic tweezer developed
here in this thesis can successfully and repeatedly apply tension to single molecules of
DNA.
51
Figure 2.5: With our tweezer, I can subject DNA to various forces over time. I can move
from low- (<0.1pN) to high (>50pN) force and back again with minimal hysteresis. This
figure shows that when moved at a rate of 0.320µm/s, DNA is exposed to increasing or
decreasing forces in very near equilibrium conditions. From this figure and the preceding
figures, one can see that our load rate is low enough to very nearly match the wormlike
chain model shown here as a dotted line. In this figure, the x-axis starts at 10µm.
Chapter 3
Native Nucleosomes under tension
In this chapter, we describe single molecule experiments on arrays of
reconstituted native nucleosomes. In addition to the steps needed to carry out forceextension measurements on single DNA molecules, these experiments require methods
for purification of native histones and a technique for reconstituting nucleosomes on an
extended DNA tether, We describe our approach to these problems here and in associated
appendices. This is followed by a section describing our results and then a section
discussing the implications of our findings for native nucleosome stability and force
response. Portions of this chapter will be submitted in manuscript form to be considered
for publication in a research journal.
3.1 Instrumentation & Technique
3.1.1 Introductory Material
3.1.1a Magnetic Tweezers
All experiments were performed in the magnetic tweezers described in chapter 2
to allow visualization of changes in a DNA-tethered pair’s end-to-end extension. The
only major addition to the setup described in chapter 2 is the protein injection pipette
used to introduce histones to DNA under tension.
52
53
3.1.1b Histone purification
The histone core complex was purified from cells using column chromatography.
To assess purity and enrichment of the core complex, the crude histone preparation (CH),
column flow through (FT), wash (W) and the final column eluate (E) were analyzed on
Coomassie Blue-stained SDS-PAGE gels. As shown in Figure 1, the core complex was
efficiently purified from both the cultured WIF-B cells as seen by the robust staining of
the histone subunits in the column eluate. More information regarding the purification of
histones can be found in the appendices VI and VII.
3.1.1c Histone experiments
Several more steps are needed to perform a DNA-histone interaction experiment
than those required to perform a DNA force-extension experiment. First, a second pipette
needs to be pre-loaded with a 7.5μL sample of the histone solution. Second, the second
pipette with a larger opening needs to be fixed at an angle relative to the aspiration
pipette to introduce DNA-binding proteins on demand onto the extended DNA molecule.
Third, once a DNA tethered bead pair is found, it needs to be positioned appropriately in
the magnetic field gradient to keep DNA nearly fully extended at a force below the
expected critical force of ~1.6pN.
54
The protein injection pipette was formed using a vertical micropipette puller
(KOPF Model 720) that slowly drew out a long (~2.5cm) tapper on the standard glass
capillaries mentioned above. The resulting tip had an inner diameter of ~ 15 microns. The
long tapper ensured micropipette flexibility; short (~0.5cm) tips with the same inner
diameter were more prone to fracture.
Each buffer-filled micropipette was connected to a buffer-filled syringe using
1mm-inner diameter tubing (Tygon tubing). The syringes were mounted to individual,
New Era syringe pump model NE-1000s. One pump created back pressure for the
aspiration micropipette; the other allowed spraying of the protein solution. For aspiration,
the syringe pump was set to a withdrawal rate of 5.5mL/hr – 7.5mL/hr which created a
sufficient back-pressure at the mouth of the pipette to hold the poly bead in place.
Proteins were sprayed at a rate of 1.5mL/hour. The flow was set to minimally disturb the
DNA-bead construct. See appendix V for more information on pipette manufacturing.
The aspiration pipette was loaded with Tris-EDTA buffer using a spinal tap
needle (Gerber Spinal-tap needle). Using micro-loader pipette tips (Eppendorf
microloader), I loaded the protein injection micropipette with 7.5μL of histone solution
followed by standard Tris-EDTA buffer. An air gap between the buffer and the protein
solution in the micropipette prevented mixing and contamination of the histone solution.
Details are given in appendices IV and V.
55
Figure 3.1: This figure shows that the core histone complexes are efficiently purified
from cultured WIF-B cells. Confluent monolayers of WIF-B cells were harvested and
histones purified.
The proteins in the preparative samples and final eluates were
separated by electrophoresis and stained with Coomassie Blue. The molecular weight
standards are indicated on the left of the gel in kDa and the core histones are indicated on
the right with arrows. CH, crude histones; FT, flow through; W, wash, E column eluate.
This figure and caption are provided courtesy of Dr. P. Tuma in the Department of
Biology.
56
3.2 Results for Pulling on Nucleosomal Arrays
3.2.1 Nucleosome Array Formation and Force-Induced Dissociation
Aspiration and protein injection micropipettes were loaded as described above,
mounted via a clamp on their micromanipulator, and positioned above the objective
inside the chamber. Tris –EDTA buffer was introduced to the chamber and any air at the
tips of either pipette was evacuated. A slight back-pressure was then created in the
aspiration tip to maintain the ability to catch pairs and in the protein pipette to prevent
flooding the sample cell with proteins prior to an experiment.
The DNA tethered pairs were then gently pipetted into the chamber and the
second cover-slip was placed on the chamber. I then proceeded to find tethered pairs as
described previously.
After a tethered bead pair was captured, its response to a force of 10pN was
measured to verify that I have a single molecule of DNA. If it showed full or nearly full
extension of 16.4μm, it was brought to a low force (~1pN) region 2000μm from the
magnet. I introduced protein within this force range because at forces <0.5pN the DNA
was prone to folding and kinking, expected in the DNA entropic response (Chapter 2
Figure 2.5), masking possible binding sites and making observation of the binding and
therefore the formation of the nucleosomes much more difficult.
Protein introduction resulted in a deflection of the tethered pair from the preestablished line of force. Binding of proteins was detected by the visible shortening –
usually full compaction - of the tethered DNA.
57
Once compaction occurred, I prepared to step the magnet closer to the tethered
pair, slowly increasing the force on the nucleosomal array. First, I allowed the condensed
DNA to remain at the low-force, protein injection position for 1 minute. Then, I began to
slowly (0.320μm/s) decrease the distance between the magnet and the DNA tethered bead
pair. This slow approach corresponds to a loading rate of ~0.008pN/s from 2000μm to ~
1000μm with the force increasing from ~0.8pN to ~ 9pN. It takes 52min to travel
1000μm so over 52 minutes I increase the force acting on the DNA-histone complex by
~8pN. The force may thus be considered roughly constant for each 30sec time segment
and increasing by ~0.25pN between successive intervals. This allows us to study the
stability of nucleosomes at various forces simply by selecting different short time
segments from an extension time-trace. The experiment is continued past 1000μm until
the starting extension is recovered. A typical experiment lasts for ~2 hours.
3.2.2 Nucleosomal Arrays Under Tension
Figure 3.2 shows a typical time trace of the end-to-end extension of a captured
DNA tethered pair during the course of a DNA-protein interaction experiment. The
coarse resolution of the two axes was chosen so that data for the entire course of the
experiment could be displayed. Looking at the figure from left to right, we see DNA
subject to a low force of ~0.8pN, with ~12μm extension, just prior to the introduction of
histones. I then introduce the histones at this low force position 2000μm from the magnet
using the protein micro-spray technique described earlier. The sharp drop-off in extension
58
that follows shows the rapid nature of the histone binding. I find that histone binding
followed by nucleosome formation occurs in a rapid manner, taking less than one minute.
Typically, I observed 90-95% contraction happening within 25sec. This prevented us
from observing an individual nucleosome form under most circumstances.
After contraction, the compacted tether is kept under constant tension for 1min;
arrow 1 in Fig. 3.2 marks the end of this 1 minute interval. Between points marked by
arrows 1 and 2, the force is slowly ramped up reaching ~1.6pN at arrow 2. From
theoretical arguments, nucleosomes under physiological conditions are expected to be
stable at forces below the critical force of ~1.6pN, with no net change in <z> due to
dissolution of nucleosomes. Nevertheless, some re-extension of the tether in the form of
nucleosomal ruptures is typically observed even in this force range. In this experiment,
<z> increases somewhat more rapidly than is usually the case. Below 1.6pN, I should in
principle be able to observe equilibrium nucleosome on-off fluctuations. I have some
evidence in our low-force data for these fluctuations but I currently lack the
computational ability to discern fluctuations in this force range.
Arrows 2 and 3 in figure 3.2 demarcate the region where the tether is subject to
forces between ~1.6pN and ~12.5pN. Re-extension from ~1.5μm at 1.6pN to ~8μm at
~14pN is roughly linear over time with a small slope. In fact, <z> changes in a step-like
manner with jumps of integral multiples of ~50 nm indicating that the release of one or
more nucleosomes is the primary driver of tether re-extension. The steps are not visible
on the resolution chosen for the extension axis in this graph; see discussion below for
more on the data regarding the individual steps. I also determine the typical time between
59
ruptures and find it to be between ~ 10 to ~ 15s (see the section on step-duration
histogram.)
The time interval defined by arrows 3 and 4 shows how the tether responds to
forces between ~13pN – ~20pN, when it finally recovers full extension. Nucleosomes
are much less stable for these large forces, which is apparent in the sharp increase in
extension during this time. It is also seen in the reduction in duration between successive
ruptures from ~10 s to fractions of a second.
Further step-like events are possible even after the starting extension has been
reached (and for correspondingly higher forces.) This is because the tether is not fully
extended to begin with, which means that some nucleosomes can stabilize DNA loops
without producing a reduction in <z>. I test for this by continuing to extend the tether
beyond ~12
m to locate further unfolding steps. However, I generally find that this
rarely happens, as I would expect since the tether is nearly fully extended at ~1pN, the
applied DNA tension when I spray with histones.
All our force-extension measurements on histone-compacted tethers show an
extension plateau for forces below ~1.6pN (between arrows 1 and 2) which I refer to as
the low force response, a regime where <z> changes slowly in discrete steps with interstep duration ~10 s (between arrows 2 and 3) which I refer to as the medium force
response, and a region where <z> increases rapidly with step-to-step transitions taking
place ~0.1 s (between arrows 3 and 4), which I refer to as the high force response. The
three response regimes correspond well to the following force ranges: 0pN - 2.5pN,
2.5pN - 12.5pN, and 12.5pN - 25pN, respectively.
60
.
4
3
1 2
Figure 3.2: This figure shows the time evolution of DNA end-to-end extension during a
typical DNA-histone interaction experiment. In this figure, at time zero (i.e. frame 0),
naked DNA is subject to a low 0.8pN force producing the expected DNA extension
(Chapter 2 Figure 5). Moving from left to right, histones are introduced to the buffer
chamber and the rapid protein binding and nucleosomal formation is observed as the
steep slope down labeled bi arrow (1). Next, one can observe a period of relative stability
while the nucleosomes are subject to 0.8 to 1.6pN referenced by arrows (1) to (2). The
next region, (2) to (3), highlights a long stretch of gradual extension over constantly
increasing forces from ~1.5pN to ~12.5pN. In this figure, one can observe a steep slope
up followed by a region of steadily increasing extension. The third region between arrows
(3) and (4) highlights the response of bound proteins to higher forces >12.5pN. In this
region, the sudden increase in extension over a time of ~ 30 seconds suggests that an all
or nothing threshold has been reached at which nucleosomes become very unstable.
61
A key feature of my data is that individual ~50nm jumps in extension can be
clearly distinguished at these forces. Fig. 3.3 illustrates the basic concept of how and why
we are able to observe quantized jumps in extension data. When histones bind to DNA
and form nucleosomes, each nucleosome “spools up” approximately 50nm of DNA. For
high enough forces, the nucleosomes –three are shown in the figure – can rupture
resulting in tether re-extension by increments of 50nm, in this case < z> ~ 150 nm since
three nucleosomes dissociate.
In Fig.3.4 I present a snap shot from higher resolution extension vs. time data for
compacted DNA. These data start from a force of ~1pN that is subsequently ramped up.
In this figure, the force reaches ~1.7pN, highlighting the transition from no net change in
<z> below 1.6pN to step-like unfolding of the tether just beyond the Marko-Siggia
critical force.
As the force is slowly increased to about 1.6pN, there is essentially no change in
extension but extensional fluctuations about <z> are observed with roughly equal number
of < z> ~ 50 nm and < z> ~ -50 nm events with no net increase in <z>. In this figure,
about 15 such events are seen. This is what I expect for forces below 1.6pN. The same
behavior is seen in multiple experiments and strongly suggests that nucleosomes are
forming and rupturing in a state of equilibrium. When 1.6pN is reached – as marked by
the arrow - I see <z> begin to increase in ~50 nm steps. This is the first time nucleosomes
have been shown to rupture at the predicted critical force.
I also find that ruptures accounted for an average stabilization free energy
(average over the force range for each data set and then an average over the data sets for
F
62
Force ~ 0.8pN
Histone
Octomers
Nucleosome
Formation
Nucleosome
Rupture
150nm of length lost
50nm per nucleosome
Force > 1.6pN
Figure 3.3: In this figure, I present the principle of our experiment. At the top, I show
naked DNA at low force that has been sprayed with a histone solution. The next figure
shows some proteins binding and forming nucleosomes that leads to reduced the end-toend extension of the DNA under minimal tension. The last figure of the panel shows the
behavior of the nucleosomes at forces greater than the critical force. Here the
nucleosomes are popped off and seen as steps in our time traces.
63
Figure 3.4: This figure shows a time trace of DNA extension highlighting the stability of
the reconstituted nucleosomal arrays at forces <1.5pN. This figure also highlights the
force at which the rupture events begin at ~ 1.5pN. This rupture beginning is visible on
the right side of the main figure. The inset shows the experiment from a few moments
before protein introduction, through the reclamation of all the DNA end-to-end extension
approximately 50 minutes later. The inset also highlights the specific region in the
experiment from which this step data of the main figure was drawn.
64
comparable extensions) of ~18.6 kBT per nucleosome for forces between ~1.0pN to
~2.5pN.
Figure 3.5 is selected from typical extension data covering ~2.5pN to ~12.5pN. It
shows how <z> changes due to nucleosome unbinding at ~ 6.5pN. The relative temporal
stability of the nucleosomes is also evident. Several seconds pass between rupture events.
Note the simultaneous release of two nucleosomes observed as a ~100nm jump in
extension. Here I find the average binding free energy to be ~38kBT, similar to that
observed in previous works (see discussion.) The inset marks the location in the full
extension trace from which these data originated.
Beyond 12.5pN, a rapid jump in extension is typically observed. Fig. 3.6
highlights the extensional response around ~20pN. The nucleosomal unbinding events
take place in quick succession, ~ 1 unbinding every ~0.6s, which I observe in all of our
experiments. The inset marks the location in the full extension trace from which these
data originated. As the reader can verify by comparing the insets to figures 3.6, 3.7, and
3.8, the data are drawn from three different experiments to show the repeatability of the
effects I describe here.
3.2.3 Step Size Distribution
Figure 3.7 presents an average distribution of the observed step-sizes from all 10
experiments over the entire force range scanned for each experiment. The histograms
revealed peaks showing (expected) 50nm and 90nm jumps in extension. These sharp
65
Figure 3.5: This figure presents data from a region of moderate force between ~2.5pN
and 12.5pN. Here I present data for typical unbinding events at 6.5pN. Observe the
temporal stability of the events. There is approximately ten seconds between subsequent
rupture events. This raw data also shows clearly the ~50nm step-size of each event
strongly suggesting that these are the disruption of nucleosomes. Again, the inset shows
the entire time trace of the experiment as well as the highlighted regions of protein
response.
66
Figure 3.6: This figure highlights the quick release of nucleosomes at forces > 12.5pN.
This shows disruption events occurring approximately 0.67s apart at a force of 20pN.
There is a observable temporal instability at this force compared to the lower force
presented in figure 3.4. Again, I present the entire time trace of the corresponding
experiment highlighting the three distinct force response regions.
67
peaks provide evidence for the rupture of individual nucleosomes, and correspond to the
previously observed steps in the time traces shown in Fig. 3.6-3.9. We find that the 50nm
steps account for nearly a third of all of the disruption events irrespective of the force.
After the secondary peak, there are peaks at or near 130nm, 170nm, and 210nm. These
suggest that multiple simultaneous nucleosome ruptures can and do occur.
The nearly 90-95% compaction of the DNA upon protein introduction clearly
indicates that histones have bound to the DNA since this is not observed in control
experiments when histone-free buffer is sprayed onto the DNA. In addition, I find that in
some experiments, when I analyze the compaction phase of the extension data, I can even
make out individual 50 nm extension reduction steps, which strongly suggests formation
of nucleosome by the bound histones. But the clearest evidence that I have observed
nucleosomal formation and force-induced rupturing are the distinct 50 nm steps that I
find at various forces at or beyond the critical force. Moreover, I find that the area under
the histogram accounts for all the extension released after I applied tension to the histonecomplexed DNA tether. In other words, almost all of the recovered extension is
accounted for by discrete 50 nm or 90 nm step-like extension jumps. This would not be
expected if the histones did not form nucleosomes upon interacting with the DNA.
The bin size used here was 10nm. Using smaller bin sizes, ~1nm for instance, did
not improve the interpretability of the data since I was unable to resolve a < z> this
small. On the other hand, I found that using larger bin sizes led to an artificial reduction
in precision since many events that I could discriminate in the data were lumped together.
68
50nm
90nm
130nm
170nm
210nm
Figure 3.7: This figure presents an average distribution of step-sizes. The vast majority of
steps occur as 50nm jumps in extension as shown by the high peak in the histogram. The
average number of 50nm steps for a given experiment is ~ 32. There is also a shorter
secondary peak at 90nm. This strongly suggests the disruption of multiple, individual
nucleosomes as well as the occasional disruption of two simultaneously. After the
secondary peak, there are visible peaks at or near 90nm, 130nm, 170 and 210nm. These
suggest that nucleosomes are more probable to dissociate individually rather than all at
once. The step-finder algorithm used to identify the disruptions rounds the measured
steps to a nearest multiple of 5nm. The bin –size for counting these step events was
10nm.
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3.2.4 Nucleosome Response to Force: Step-Counts and Step-Size
In fig. 3.8, I plot a histogram of number of steps as a function of force. The bin
size I use for the force is 0.1pN. Thus, the histogram tells us on average how many steps
correspond to each force during a typical experiment. Here I do not distinguish between
steps of length 50 nm or multiples of it, but as I discussed earlier most steps correspond
to 50 nm jumps. I see a clear, low force peak centered at ~ 1.5pN, indicating that many
unbinding events occur at the critical force.
In figure 3.9 I show a force-step hybrid histogram that correlates the step-size to
the force applied. In this figure, one can see that about 95% of rupture events occur
between 1pN through ~20pN with ~65% being 50nm steps jumps. High force ruptures
account for the remaining 5% of all events and are distributed among the 50nm, 90nm
and 135nm events in the same percentages as jumps for lower forces. Surprisingly, I find
that no matter what the applied tension, single nucleosome dissociations producing < z>
≈ 50nm are the dominant type of event, although fewer jumps take place at high forces.
Figure 3.8 shows the proportion of 50nm and 90 nm step events among all steps for each
0.1pN force bin. This allows us to determine which sort of step size dominates at
different forces for a typical experiment. It should be noted that multiple, simultaneous
disruption events show no clear propensity to occur at higher forces. Thus I find that
there is no distinct correlation between step-size and force.
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~1.5pN
~2.5pN
~4.5pN
~7.5pN
~15pN
Figure 3.8: This figure shows an average distribution of step-events as a function of
force. In this figure, the data presented show the low force (~1.5pN) response of the
native nucleosomes. Many unbinding events occur at or above this critical force. A
gradual decrease in events can be explained two ways: first, few nucleosomes remain
when the arrays reach high forces, >12.5pN, nearly 50 minutes after they formed, second,
the nucleosomes have a propensity to pop off sooner under lower forces. The disruption
events occurring at or near 1.5pN correspond very well with the predicted critical force.
71
90nm Peak
130nm Peak
Increasing Force (pN)
50nm Peak
170nm Peak
Figure 3.9: In this figure, step-size, step-count and step-force data are all plotted together
to highlight the distribution of each step event to various forces. One can see that there is
no direct correlation between step-size, i.e. number of simultaneous nucleosome
disruptions and force applied. Multiple nucleosome ruptures occur throughout the force
range from 0pN to 20pN.
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3.3 Discussion
3.3.1 Advantages of this System Over Other Systems
I have developed a low-noise, magnetic tweezers system that is capable of
applying tension in the focal plane. With it, I was able to directly observe the binding of
histones to an extended DNA molecule and subsequent rupture of formed nucleosomes at
low forces where they were not detected before. Using comparatively simple particle
tracking software, I detected 50nm steps allowing us to follow the unbinding of
individual nucleosomes (and in some cases their formation).
An attribute of our design is the ability to slowly change the force acting on the
re-constituted nucleosomal arrays. Between distances of 3000µm - 2000µm from the
magnet the force increases slowly and linearly with step rates of 0.320μm/s resulting in a
force loading rate of ~ 0.008pN/s. This ensures that experiments are performed in near
equilibrium conditions. The combination of quasi-static force application and minimal
mechanical noise enables us to observe DNA-histone interactions and modulate their
binding by applied tension, especially at very low forces.
A factor that limits the number of experiments I can perform is my ability to find
tethered pairs at the bottom of the chamber. This is greatly mitigated by the design of my
aspiration system, which allows for rapid capture and release of beads, and the ability to
scan the floor of the chamber. Together, this combination typically allows me to
investigate a large number of putative tethered beads in reasonably rapid succession thus
increasing the chances that a usable DNA-bead construct will be found. Once such a pair
73
is found, I can hold on to it quite stably, enabling tethered bead pairs to be stepped into or
away from the magnet at a very slow speed.
Optical trapping techniques used in the investigation of nucleosome formation
and rupture under force have provided much information but have been limited by nonequilibrium experimental conditions (Cui and Bustamante 2000, Bennink et al 2001,
Brower-Toland et al 2002). These techniques provided the required spatial resolution to
observe individual nucleosomal ruptures but lacked the ability to modulate the forces
slowly enough to provide near equilibrium conditions when observing nucleosome
ruptures in response to force.
Vertical magnetic tweezers are resolution limited by the need to continually
refocus on a bead moving in and out of the focal plane. This hampers the ability to
resolve 50nm changes in extension. Other transverse magnetic tweezers systems have
been developed by Yan et al, Sun et al, and Chen et al. Yan et al had low-force extension
measurement limitations because they used an open flow cell design which led to
mechanical noise, buffer evaporation, and other issues (Yan et al, 2007, Chen et al 2011).
The approach used by Chen et al and Sun et al prevented them from applying a wide
range of forces since they could not bring the magnet arbitrarily close to the
superparamagnetic bead (Chen et al, 2007, Sun et al, 2008). In addition, their ability to
resolve small < z> was limited by the fact that they measured < z> from the centroid of
their bead to an arbitrary point on a glass surface, rather than as the separation between
two beads.
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3.3.2 Low Force Response: Theoretically-Predicted Critical Force Observed
After the injection of histones into the buffer chamber, upon histone binding I
always observed a rapid compaction of the DNA. Compaction of DNA typically took less
than 1 minute, often occurring as fast as 30 seconds. Under the low force of ~ 1pN and
relatively high site specific protein concentrations, I observed nucleosome formations
that behaved similarly to those reported by several other groups (Leuba et al, 2003, Yan
et al, 2007). In Leuba et al, they observed the binding characteristics of histones and
subsequent formation of nucleosomes as a function of applied force (Leuba et al 2003).
They reported significantly slower binding and formation rates while applying both
magnetic and flow-based forces to the DNA that resulted in forces of between ~3pN and
~10pN. Those forces are significantly higher than the ~1pN I applied to DNA when
observing binding of histones and nucleosome formation. In Yan et al, they report
observing rapid formation of nucleosomes under low forces but do not present detailed
data (Yan et al, 2007).
After allowing nucleosomes to form on the extended DNA tether, I slowly
scanned the force from about 1pN through the critical force f * where nucleosome on- and
off-states are equiprobable, and then on to larger forces. As I show in our results, there is
a clear transition at f * with no net change in <z> for f < f * but <z> increasing for f > f
*
with a staircase-like profile with the steps of length ~50 nm or multiples thereof. Our
results verify the predicted value of f
nucleosome stability under tension.
*
based on a statistical-mechanical analysis of
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In a theoretical study of DNA-protein interactions, Marko and Siggia addressed
the problem of stability of nucleosomes under applied tension. (Mark and Siggia, 1997).
Starting with the free energy difference of bound and disrupted nucleosomes,
F
k B T log
fl  (4)
where k b is the Boltzmann constant, T is the absolute temperature,
of protein in solution,
is the concentration
is the binding enthalpy, f is the force acting on the molecule
and l is the length of DNA wrapped around the protein, they proceeded to derive the
equilibrium force f* = 1.6pN needed to mechanically-disrupt nucleosomes..
A number of single molecule approaches have been developed to study how
nucleosomes behave under force. Nucleosomal arrays formed on DNA – the approach
used here – or chromatin fibers have been used for these micromechanical studies. Until
now, no one has provided clear evidence that nucleosomes readily dissociate at the
predicted f*, but evidence of nucleosome disruption events at higher forces has been
found. The first study along these lines was by Cui and Bustamante, (Cui and
Bustamante, 2001) who used a chromatin fibers extracted from chicken erythrocytes that
were manipulated using a combination of micropipette aspiration and optical trapping.
Although no nucleosomal disruption data were presented, they did map out the forceextension response of chromatin.
In Bennink et al, single chromatin fibers reconstituted using Xenopus laevis egg
extract were studied under applied tension, and the unfolding of individual nucleosomes
was observed, but the corresponding forces were in the range of 20pN to 40pN, an orderof-magnitude larger than what was theoretically predicted (Bennick et al, 2001). This
76
work was the first of its kind to present individual nucleosomal ruptures, but did not show
good agreement with the Marko-Siggia f* prediction. One possibility was that their
experiment was not well suited to measure f*, which was established for the condition
when force transduction was nearly reversible since the experimental design based on a
laser tweezer resulted in a force loading rate of ~38pN/s.
Brower-Toland et al reported on experiments performed using an optical trap to
study the rupture of nucleosomal arrays along a short (3684bp) segment of DNA
(Brower-Toland et al 2002). From the description of the experiment, I have not been able
to determine the force loading rate appropriate for their study. They present results of
individual nucleosomal disruptions at forces of ~15pN but do not provide data on the
stability of nucleosomes at low forces (<5pN).
Yan et al used a combination of vertical and horizontal magnetic tweezers to
study reconstituted chromatin fibers (Yan et al 2007). They present data on the formation
and subsequent extension of chromatin fibers at forces near 1.6pN, but do not report
direct observation of individual nucleosomal ruptures at this force. However, they do find
disruption of nucleosomes at forces of 4.5pN. Although their micromanipulation
approach led to low load rates – a major improvement over previous efforts – relatively
large mechanical noise that resulted from using an open experimental cell possibly
prevented clear observation of nucleosome disruption events at the predicted f*.
The main reason why single molecule nucleosome stability assays using optical
traps did not observe nucleosome dissociation at the predicted f* was the very high
loading rates involved which resulted in non-equilibrium conditions. (Bennink et al,
77
2001, Brower-Toland et al, 2002, Pope et al, 2005). On the other hand, in previous
magnetic tweezers studies, the loading rate problem was solved but the limiting factors
were data acquisition rates of vertical tweezers and high noise levels associated with
open-sample-cell transverse magnetic tweezers (Yan et al 2007).
In our experimental setup, I quasi-statically change the amount of force applied to
the reconstituted nucleosomal arrays. The histones initially bind to DNA at a very low ~
0.8pN force. The compacted DNA is then subject to a slowly changing force (0.008pN/s).
Although this is not in pure thermodynamic equilibrium, I still observe the transition
from stable nucleosomes below ~ 1.5pN to discrete 50nm jumps in the DNA’s end-to-end
extension above this force, thus confirming theoretically-predicted value for this force.
By using a semi-closed cell with a glass slide “roof,” mechanical noise is reduced by
several orders of magnitude, allowing us to collect low-noise, high-resolution extension
data to measure 50nm steps at the critical force. I also nearly eliminate any evaporative
currents, one reason the experiments can last 2 – 3 hours
I find that in most of our data sets the transition force f* ≈1.6pN, but there is a
distribution in f* confined to a narrow range from 1.1pN to 1.7pN. I even find some cases
where f*≈ 1pN, i.e. the tether compacts after the protein micro-spray, and upon
compaction re-extends almost immediately at the same force, with the rest of the forceextension behavior that is identical with other experiments. But for forces below 1pN
there is no net increase in <z> in any of our experiments.
78
3.3.3 Moderate and High Force Response
Brower-Toland et al present fixed-force nucleosome disruption data showing a
rapid chain of quantized events of ~25nm when nucleosomes are exposed,
instantaneously, to forces of 17.6pN and 20.2pN; multiple nucleosomes are found to
rupture each second (Brower-Toland et al 2002). Although I do not find distinct 25nm
extension release events, our high force data presented here does reproduce their
observation of rapid nucleosomes disruptions. For instance, in figure 2, I observe 3
nucleosomes rupturing within ~1.5s. They also find that the initial rapid release of
nucleosomes is followed by period of less frequent ruptures. This feature is absent in my
force-extension data most likely due to differences in experimental conditions, most
importantly the near-equilibrium force loading method I have used. Nevertheless, at high
forces, I do observe the rapid release of several nucleosomes.
I also recapitulate Bennink et al’s report of nucleosomal ruptures as a sudden
release, an “all-or-nothing event” in which all of the bound nucleosomes dissociate nearly
simultaneously Bennink et al, 2001). They attribute this behavior to the high loading rate
of 38pN/s used for the experiment, but our results suggest – see fig. 8 - that this effect
may be intrinsic to the organization of nucleosomes and could indicate the presence of a
cooperative binding effect.
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3.3.4 Free Energy vs. Force
I also determine the average nucleosome stabilization free energy F over 0.5pN
to 2.5pN and find that F ≈ 18kBT. The method used to calculate this can be found in
Appendix XII. This is less than ~ 20k BT used by Marko and Siggia and reported by Yan
et al and Brower-Toland et al. (Cui and Bustamante and Bennink et al only report high
force data and do not report on any free-energy calculations.)
In Bennink et al the high force ruptures correspond to
F of ~ 250k B T . They
addressed their inability to make low force measurements and attributed that limitation to
the 38pN/s loading rate inherent to their velocity clamp (Bennink et al, 2001).BrowerToland et al calculate a F between ~ 36.0k BT and 38.0k BT for the release of short 40bp
segments of DNA from a nucleosome at forces of either 17.6pN or 20.2pN (BrowerToland et al, 2002). This would suggest a free-energy requirement of ~ 135.0k BT for total
dissociation. The difference between their results and ours is quite stark. Bennink et al
admit that their high 38pN/s loading rate probably drove the nucleosomes out of
equilibrium; as discussed earlier, that was also probably the case with the experiments of
Brower-Toland et al. Thus, the discrepancy between their results and mine can be
attributed to the difference in experimental methods used.
Pope et al found that the loading rates variability from experiment to experiment
was responsible for the differences in the amount of free energy required to dissociate
some or all nucleosomes (Pope et al 2005). They reported an inverse relation of loading
rate to energy with low loading rates requiring more work, 25.0k BT , and high loading
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rates requiring less work 20.0k b T . The range of loading rates spanned 3.5pN/s to
560pN/s. Although I find a lower free energy barrier of ~ 18.6k B T , I also present data that
shows ~10% of observed unbinding events required ~ 25.0k BT . One way to reconcile my
results with those of Pope et al is to posit at least two distinct subpopulations of
nucleosomes with different stabilization free energies. In addition, it is also possible that
only at very low loading rates will nucleosomes with the smaller F pop-off as a distinct
group. Thus, if the force is applied too rapidly, these nucleosomes will dissociate but at a
larger force thus leading to the higher F = 25.0k BT . Therefore, a high loading rate may
make it impossible to distinguish between these sub-groups using applied tension.
3.3.5 Step-Size and Applied Force
Yan et al report the behavior of nucleosomes subject to forces of 4.5pN, 9.6pN
and 15pN (Yan et al, 2007). Their data for 4.5pN and 9.6pN agrees well with ours for the
force range shown in Fig. 8. I observe nucleosomal stability similar to theirs with one
noticeable difference: the time between disruption events. In my study, I observe ~ 15s to
25s between events where they observe 30s to 45s between events. This is most likely a
result of the different methods used to form nucleosomes. In my work, I introduced only
native, octomer-core histones; in their work, they introduced Xenopus extract which
contained not only the core histones but also the linker histones and other DNA binding
proteins.
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For high forces, Bennink et al observed much larger 65nm steps (Bennink et al,
2001). I do not find evidence for these jumps in our data. This may again be a result of
having used Xenopus egg extract and the high 38pN/s loading rate and in line with their
observations of a longer time between events.
Brower-Toland et al claim that in the low force regions, nucleosomes are
gradually peeled off in a smooth manner (Brower-Toland et al, 2002). Their data show
that when exposed instantaneously to forces of 17.6pN and 20.2pN, multiple
nucleosomes rupture each second. These fixed-force nucleosome disruption data show a
rapid chain of quantized events of ~25nm. Our high force data presented here shows
good agreement with this behavior with the notable exception being the step-size. I
consistently observed the 50nm peak while their data suggests that most events occur as
small 25nm changes in extension. Again, I attribute this behavior to the optical traps used
and the high loading rates acting on DNA and nucleosomes.
Pope et al present step-size distribution data with distinct peaks at ~30nm and
~60nm (Pope et al, 2005). They present two sets of histograms: one set that shows step
distributions from 0 – 50pN and the other set that shows step-size distributions for force
windows of 0pN – 20pN and 20pN to 40pN. In these data, they have shown few rupture
events occurring in the force range of 0pN to 20pN. Although they present data showing
disruption events at forces less than 10pN, the lack of data showing quantized events
suggests, again, difficulty in resolving low force extension jumps when using optical
traps. Their data does not provide any insight into the rate at which nucleosomes rupture
as a function of force.
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In experiments with optical traps (Pope et al, 2005, Bennink et al, 2001, BrowerToland et al, 2002), the step-size distributions at various forces showed peaks at ~30nm
and ~60nm. In our study and in the work of Yan et al, a peak was observed at ~50nm. In
all of these experiments, nucleosomes were formed using either Xenopus egg extract or
purified histones. Neither our work nor the work of Yan et al presents data that shows a
distinct and significant peak at 30nm. Both do show secondary step events roughly
corresponding to 90nm, 140nm and 180nm. I believe this to be due to the low loading
rates achieved with the magnetic tweezers.
Chein and van Noort also highlight the effect that experimental design can have
on studies of nucleosomal unbinding events (Chein and van Noort, 2009). For example,
the primary peaks at ~30nm found in some studies may be due to partial unwrapping of
DNA from the nucleosome caused by higher loading rates associated with optical traps
(Kruithof et al, 2009). This suggests that the low loading rates associated with magnetic
tweezers may provide a more stable platform for observing tension-modulated
nucleosome formation and dissolution.
3.3.6 Step Size Does Not Depend on Force
Throughout the force range, I observe no significant correlation between step-size
and force. This is in contrast to several earlier works by Bennink et al and Pope et al. I
believe that the low loading rate that I use is responsible for this observation. Fig. 8
clearly shows the distribution of rupture events through the force range. In it there is no
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observable bias suggesting that multiple simultaneous ruptures occur at higher force. On
the contrary, I show that there are single, double, and even triple rupture events occurring
throughout the entire force range.
This is in contrast to the observations made in other works of Bennink et al and
Bower-Toland et al but, as discussed above, these experiments used optical traps with
much higher loading rates. If there is an all or nothing response, it should be observed as
a clean break in the extension vs. time curve. As I observe in figures 4-6, this is not the
case. These time traces show no sudden, instantaneous, clean jump in extension. Fig. 8
shows that there is no significant relation between step-size and force ranges. I conclude
that this is a result of the near equilibrium conditions under which tension was applied to
the compacted DNA from ~ 1pN to 20pN of force. I also believe that the high resolution
imaging allowed us to observe all of the individual jumps.
Chapter 4
Native vs. Post-Translationally Modified Nucleosomes Under Tension
In this chapter, we describe single molecule experiments on arrays of
reconstituted post-translationally-modified (PTM) nucleosomes, specifically hyperacetylated nucleosomes. After a brief discussion of some significant aspects of histone
acetylation, we describe here and in the appendix the additional steps involved in
acetylating histones and reconstituting nucleosomes from them. This is followed by a
section describing our results and a detailed comparison of the behavior of native and
PTM nucleosomes under applied tension. This chapter concludes with discussion of the
implications of the findings. Portions of this chapter will be submitted in manuscript form
to be considered for publication in a research journal.
4.1 Post-Translational-Modifications: Structure and Significance
The post-translational-modification (PTM) of a protein is the subject of intense
research not only because of the many forms they take but also because they play a
critical role in the in structure and function of many proteins. Acetylation, methylation,
phosphorylation and ubiquitination were all mentioned in chapter 1 as types of PTMs that
can happen to a protein. These PTMs are the result of an additional chemical group to a
residue amino acid of a protein. Typically, these amino acids lie along side chains, or
tails, of the proteins. Other types of PTMs can occur – various folding conformations that
do not involve the addition of chemical groups, but they are not discussed here.
85
85
In this work, I address the effect of having all or nearly all of the lysine residues
in the core histones –H2A, H2B, H3 and H4- acetylated, so-called hyper-acetylated
histones. The histones that comprise the core of a nucleosome structure all have rather
long amino acid residue tails composed predominantly of lysines that can be acetylated.
Histones and DNA interact through electrostatic attraction. Core histones have a
net positive charge which leads to strong electrostatic attraction with DNA, strong
enough to wrap a stiff 50nm-long segment of DNA – 50nm is DNA’s persistence length around them. The addition of acetyl groups acts to neutralize the positive charge in the
core histones (Hansen, 2002). This charge neutralization then lessens the attraction
between the core particle and the DNA. This reduction in attraction significantly weakens
the attractive forces holding together the nucleosome particle, making it both more
susceptible to popping off under external tension and more accessible to being opened by
internal molecular machines (Hansen, 2002).
4.2 Comparison Study of Native and Post-Translationally Modified Histones
4.2.1 Protein Introduction & Re-extension.
The procedure for introducing hyper-acetylated histones is the same as that
described in chapter 3 where the introduction of native histones is discussed. The
protocol for producing hyper-acetylated histones is in appendix VII.
In all of the experiments performed, I introduced histones, native or modified, at a
low force of ~1pN. Figure 4.1 provides a diagram of the principle behind a typical
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experiment. Initially, the DNA at ~ 1pN is not fully extended and is not taught. Rather,
the DNA is fluctuating in the thermal buffer bath in the sample cell with a small amount
of tension applied by our bar magnet 2000μm away. I introduce the histones at this force
because I need to observe the binding of these proteins to the DNA – at lower forces,
DNA is shortened by thermal kinks and bends making observation of binding difficult, at
higher forces the tension will prevent binding. After introduction, I can observe the
shortening of the DNA under tension as histones bind to and form nucleosomes. In the
figure I show three nucleosome formations leading to the loss of ~150nm in extension.
After the histones have bound and formed nucleosomal arrays, I subject the nucleosomal
arrays to increasing force. As the tension increases, I can observe the quantized changes
in DNA’s end-to-end extension corresponding to the rupture of individual nucleosomes.
In a typical native histone experiment represented by Fig. 4.2A, I observe three
distinct nucleosomal responses to force: low force response to 2.5pN or less, moderate
force response to forces between 2.5pN and 12.5pN and a high force response to anything
greater than 12.5pN. In a typical hyper-acetylated histone experiment represented by Fig
4.2B, I do not observe the same type of behavior. Rather, I typically observe a more rapid
dissociation of nucleosomes under low forces typically less than 5pN.
Looking at figure 4.2A from left to right, we see DNA subject to a low force of
~0.8pN, with ~12μm extension, just prior to the introduction of histones. I then introduce
the histones at this low force position 2000μm from the magnet using the protein microspray technique described earlier (see chapter 3 section 1.3). The sharp drop-off in
extension that follows shows the rapid nature of the histone binding. I found that histone
87
Force ~ 0.8pN
Histone
Octomers
Nucleosome
Formation
Nucleosome
Rupture
150nm of length lost
50nm per nucleosome
Force > 1.6pN
Figure 4.1: In this figure, also seen in chapter 2, a sketch of the principle of the
experiment at the various stages is shown in the main figure. Loose DNA at low force is
sprayed with a protein solution. Some proteins bind and form nucleosomes and reduce
the end-to-end extension of the DNA under minimal tension. At forces greater than the
critical force, the nucleosomes are popped off and seen as steps in our extension-time
traces.
88
binding followed by nucleosome formation occur in a rapid manner, taking less than one
minute. Typically, I observed 90-95% contraction happening within 25sec. This
prevented me from observing individual nucleosomes form under most circumstances.
Looking at figure 4.2B from left to right, we again see DNA subject to 0.8pN of
force with an extension of ~13μm prior to the introduction of hyper-acetylated histones.
We again observe the rapid, ~1min, binding of histones and formation of nucleosomes as
the sharp drop-off in extension. I typically observe the same level - 90-95% - of DNA
compaction seen in the native histone experiments.
After contraction, the compacted tether is kept under constant tension for 1min;
arrow 1 in Fig. 4.2A and 4.2B marks the end of this 1 minute interval. Between points
marked by arrows 1 and 2, the force is slowly ramped up reaching ~1.6pN at arrow 2.
From theoretical arguments, nucleosomes under physiological conditions are expected to
be stable at forces below the critical force of ~1.6pN, with no net change in <z> due to
dissolution of nucleosomes.
I begin to see differences in the dissociation of native and hyper-acetylated
nucleosomes in the force region of ~0.5pN to 12.5pN. In all of the native histone
experiments, I observed 1301 nucleosome rupture events of various quantized step sizes.
Aggregating the data from 10 hyper-acetylated histone experiments, I observed 842
nucleosome rupture events. In the native histone experiments, 646 nucleosomes rupture
events occurred when subject to forces < 2.5pN representing ~50% of all unbinding
events. In the hyper-acetylated experiments, 661 nucleosomes ruptured in the same force
89
A
4
3
1
2
3
1
B
2
90
Figure 4.2A and 4.2B: 4.2A shows typical native histone experiment time trace
highlighting three distinct regions of nucleosome force response. In this figure, naked
DNA is initially subject to a low 0.8pN force and shows the corresponding extension
expected of DNA subject to this low force. Moving from left to right, the histones are
introduced to the buffer chamber and their rapid binding to DNA is observed as the steep
slope down in the figure. Next, from arrow 1 to arrow 2, one can observe a moment of
relative stability while I let the DNA-protein structure rest at the low force. Between
arrows 2 and 3, I highlight a long stretch of gradual extension over forces from ~1.5pN to
~12.5pN. Between arrows 3 and 4, I highlight the response of bound proteins to forces
>12.5pN. In this region, the sudden increase in extension over a time of ~ 30 seconds
suggests that an all or nothing threshold has been reached at which native nucleosomes
become very unstable. Figure 4.2B: This figure shows time trace of a typical PTM
histone experiment in which I highlight the lack of distinct regions of force response.
PTM histone experiments typically show a release of histones more rapidly than native
histones. Arrow 1 indicates the rapid formation of nucleosomes and arrow 2 shows the
rapid dissociation of the nucleosomes at a force of ~ 2pN for this particular experiment.
This curve shows, at arrow 3, that upon reaching a force of ~1.6pN, nearly all of the
nucleosomes dissociated over a time of ~1minute.
91
range representing ~79% of all unbinding events. For unbinding events occurring
between 2.5pN and 12.5pN, the differences are much more telling with ~42% of native
~18% of hyper-acetylated nucleosomes unbinding. In both sets of experiments, few
events occurred at forces greater than 12.5pN with ~ 8% and 3% for native and hyperacetylated, respectively. Thus hyper-acetylated experiments show no distinct forceresponse regions comparable to those observed in the native histone experiments, the
hyper-acetylated experiments showing more rapid dissociation of nucleosomes after
compaction as illustrated in Fig. 4.2B. (One hyper-acetylated experiment proved to be the
exception with few hyper-acetylated histone nucleosomes lasting long enough into a
typical hour long experiment to rupture at forces greater than 12.5pN.)
4.2.2 Step Histogram
Figures 4.3A (reproduced from chapter 3) and 4.3B are histograms showing the
average step-sizes observed in native and PTM experiments, respectively. Figure 3B was
produced using the same algorithm and bin size as figure 3A – 10nm (see also discussion
in chapter 3 for this choice of bin size). These histograms show the average of all events
encompassing the entire range of applied forces. The histograms revealed peaks showing
(expected) 50nm and 90nm jumps in extension. These sharp peaks provide evidence for
the rupture of individual nucleosomes. In Fig 4.3A, I present data showing that 50nm
steps account for nearly half of all of the native disruption events irrespective of the
force. After the second peak, there are peaks at or near 130nm, 170nm, and 210nm. These
92
A
50nm
90nm
130nm
170nm
210nm
B
50nm
90nm
130nm
170nm
210nm
Figures 4.3A and 4.3B: These two histograms show the differences in the unbinding of
native and PTM nucleosomes as a function of rupture size. Figure 3A, also seen in
chapter 3, shows a distinct 50nm peak strongly suggesting that nucleosomes formed from
native histones rupture in single events. Figure 4.3B shows that nucleosomes formed
from PTM histones have a broader step-size distribution. Many more 90nm and 130nm
events occur with modified nucleosomes than with native. This suggests that the more
instable PTM nucleosomes are more susceptible to simultaneous rupture events.
93
suggest that multiple, simultaneous nucleosome ruptures of 2, 3, 4 or more can and do
occur. In Fig. 4.3B, I present data that shows an observable difference in the distribution
of step-sizes. The data shows that hyper-acetylated histones have a more pronounced
secondary peak at 90nm as well as more multiple disruption events than the native
histones with about one third of all events being single jumps of 50nm and a quarter
being two simultaneous jumps releasing two nucleosomes.
4.2.3 Force Histogram & Force-Step hybrid Histogram
In figures 4.4A (reproduced from chapter 3) and 4.4B, I show histograms of the
average number of steps as a function of force, native (Fig 4.4A) vs. hyper-acetylated
(Fig 4.4B). (Figure 4.4B was produced using the same algorithm and bin size as figure
4A – 0.1pN - see also discussion in chapter 3 for this choice of bin size.) Looking at both,
the response of each type of histone is clearly discernable. Figure 4.4A shows that,
throughout the force range, native nucleosomes remain stable with many rupture events
occurring at forces of 15pN and more. On the other hand, in figure 4.4B, the sharp peak
at ~1.6pN highlights the relative instability of hyper-acetylated nucleosomes and also
shows that the majority of ruptures occur at forces < 2.5pN. In fact most of the unbinding
events occur at 1.6pN with a few outlying events between 10 and 20pN. In both sets of
experiments, we see a clear, low force peak centered at ~ 1.6pN, indicating that many
unbinding events for both types of histones occur close to the critical force indicating that
the critical force is the same for both kinds of histones.
94
~1.6pN
A
~3.0pN
~4.5pN ~6.5pN
~15pN
B
~1.5pN
~2.5pN
~4.5pN
~7.5pN
~15pN
Figures 4.4A and 4.4B: In these two histograms, we see a distinct difference in the
behavior of the native and modified histones as a function of force. The native histones,
figure 4A also seen in chapter 3, have many events occurring throughout the force range
of ~1pN to ~20pN. The unbinding events, although peaking at the low critical force of
~1.6pN, are distributed throughout the observed forces in statistically significant
amounts. Comparatively, the PTM histones are sharply peaked at ~1.6pN in figure 4B.
The unbinding events occur almost exclusively at force below 5pN. This suggests that
they are much more unstable at the higher forces that native histones can withstand.
95
In figures 4.5A and 4.5B I present the step counts, step-sizes, and the forces
associated with these events. In figures 4.3A, B and 4.4A and B I presented the number
of events per jump size and the number of events per force, respectively for native and
PTM nucleosomes. In figure 5A & B, I combine those two sets of data showing how
extension jumps - single or multiple nucleosomal disruptions - are correlated to applied
force. I also provide evidence that there is no distinct correlation between step-size and
force for both native and hyper-acetylated histones in figures 5A and 5B. Most rupture
events for both native and PTM nucleosomes occur as 50nm events throughout the wide
band of forces. As can be seen in the histograms above, small numbers of rupture events
occur at high forces. For hyper-acetylated histone nucleosomes, ~95% of all ruptures
occur in the low and intermediate force ranges from 0.5pN to ~5pN. The native histones
display far more stability throughout the same force range with many events occurring at
forces up to and including 15pN and beyond. High force ruptures account for only about
5% of all events.
When step-size, step-count and step-force are all plotted together, one can clearly
see the difference in the binding behavior of native and PTM formed nucleosomes. First,
I observe that nearly 80% of all ruptures occur at forces of ~13pN or less and that there is
no indication that multiple, simultaneous rupture events are a function of applied force.
I also find that for native histones ~ 65% of all nucleosomal ruptures occur as single
events. It is the dominant type of rupture and is populated throughout the range of forces
from ~1pN through ~20pN with fewer events happening in the high force limits. Also, I
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Increasing Force (pN)
50nm Peak
90nm Peak
130nm Peak
90nm Peak
130nm Peak
Increasing Force (pN)
50nm Peak
170nm Peak
Figures 4.5A and 4.5B: In these force-step hybrid histograms I show that there is no
correlation between applied force and step-size for either native or PTM nucleosomes.
Figure 5A, also seen in chapter 3, shows that a large number of step events occur at
forces greater than 12.5pN. Figure 5B shows the dominance of low force events for PTM
histones, as shown by the abundance of blue color in the figure representing forces less
than ~7pN.
97
expected that higher forces would lead to more simultaneous rupture events but the data
show that the proportion of single and multi step events at all forces is roughly constant.
For hyper-acetylated histones, the results are much different. In those results, one
can clearly see the predominance of low force events, shown as blue hues, and the limited
number of high force rupture events, shown in orange and red hues. The hyper-acetylated
force-step hybrid histogram shows a slight indication that higher forces resulted in more
dual-rupture events. The difference is subtle but noticeable in this comparison. This small
few % difference is indicative of the differences at low forces less than 2.5pN. The data
also show good agreement with all other previous data present which averaged over all
experiments.
4.2.4 Step-Duration Histogram
In figure 4.6, I compare the stability of each type of histone in the range of force
from 0.5pN to 12.5pN because, for both types of histones and corresponding
nucleosomes, most events occur in this range. I observe that the average time between
rupture events is nearly the same for both native and modified nucleosomes with a time
between events of ~ 5.8s for each type. The median time between ruptures tells a slightly
different story, however. The median suggests that native nucleosomes dissociate more
slowly than the modified nucleosomes. With the median time between ruptures of ~ 2.97s
for native and ~2.47s for modified, clearly more events are happening more rapidly with
the modified nucleosomes.
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4.2.5 Free-Energy Comparison
I report the first single molecule data showing free-energy differences between
nucleosome reconstituted from native and modified histones. In figure 6 I compare the
free energy needed to release nucleosomes in a force range of 0.5pN to 25pN, so that I
encompass nearly all of the rupture events. As I have indicated throughout this work,
clear differences are observable in the unbinding data, with none as critical as this. An
average of ~ 51k BT and
nucleosomes,
~ 40k BT is needed to dissociate native and modified
respectively,
implying
an
average
free
energy
difference
of
~ 11k BT between them. The median is also indicative of the differences in binding
affinities associated with native and PTM nucleosomes with destabilizion by a median
free energy of ~ 31k BT and the nucleosomes by ~ 20k BT , respectively
99
Figure 4.6: This figure shows the comparison of time between ruptures between
nucleosomes formed from native and PTM histones. The average duration of time
between events is almost identical, but the median time between events shows a subtle
but significant difference. The half second difference between the median times of native
and PTM histones shows that PTM histones have less time between rupture events than
native.
100
Average Free Energy
Native 41.4kBT
Average Free Energy
Modified 27.3kBT
Figure 4.7: This figure shows the distinct differences in the binding energetics of
nucleosomes formed from native histones and PTM histones. In all of the experiments,
across al observed forces, I show that nucleosomes formed with native histones require
far more energy to dissociate than the nucleosomes formed from PTM histones. The
differences between native and PTM, median and average energies are in close
agreement. Both the average energy and median energy show a ~ 11k BT difference in free
energy required to dissociate nucleosomes.
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4.3 Discussion
4.3.1 Comparison to Other Works
In this study, I report on the observed characteristics of nucleosomes formed
using either native of hyper-acetylated histones. I report the low, critical force rupture of
each being in complete agreement with the theoretical prediction of Marko and Siggia.
Next, I report on the similar step-size distribution for the rupture events of native and
PTM histones suggesting that all of the observed events are nucleosome disruptions.
Finally, I report on the difference in the binding kinetics between native and hyperacetylated histones.
Acetyl groups function to screen the negatively charged DNA from the positively
charged proteins has been well established in batch assay studies (Jenuwein et al, 2001,
Clapier et al, 2002, Hassan et al, 2001, Tse et al, 1998, Carruthers et al, 2000). But the
binding affinity studied here, at the single molecule level, opens the door to mapping out
the nature of individual residues contributions to the binding affinity of nucleosomes.
Here, with all possible residues acetylated, I can compare the free energy response of
native to hyper-acetylated histones.
The low noise and low loading rates of our magnetic tweezers enabled me to
detect differences in the response of the two kinds of nucleosomes to applied tension –
see chapters 2 and 3 for a detailed comparison of our approach to single DNA
micromanipulation with others.
I introduced only purified, core-histones to a DNA molecule under ~ 1pN of
tension in a 1mg/mL Tris-EDTA solution. This allows us to attribute observed
102
differences in the data between native and PTM nucleosomes to the changes made to
native histones. I then introduced either native core histones or hyper-acetylated core
histones with all or nearly all lysine residues acetylated. When observing the formation
and rupture of nucleosomes, I ensured that nucleosomes form by starting at forces <1pN
to optimize, and proceeded to increase the force until all were eventually removed up to
forces of ~25pN.
I avoided the complications of performing a chromatin re-constitution experiment
with Xenopus leavis egg extract in which multiple other proteins and small molecules, as
well as core histones, are introduced that can bind to DNA, thus obscuring the results
with possible misinterpretations.
The hyper-acetylated and native histones were used here to show a clear and
distinct difference in the binding affinities. The acetylated histones had all or nearly all of
the available amino acids in the tails acetylated. This complete or nearly complete
acetylation allowed us to create an experiment that eliminated possible sources of
confusion that can exist in experiments using reconstituted chromatin. With this all-ornothing approach, I present data that compares the behavior of native, physiological
histones, which can contain some PTM’s, with the extreme, hyper-acetylated histones in
order to show a distinct difference in binding affinity.
In Simon et al, a single molecule study using AFM, the dynamic changes in
nucleosome behavior due to the acetylation of specific lysines in the tails of histones H3
and H4 was addressed. They reported on the changes in force response of nucleosomes
due to the acetylation of residues H3 (K56), H3(K115), H3(K122), H4(K77) and
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H4(K79). They recognized the apparent structural significance of these particular
residues in the tails of H3 and H4, but do not address the roles of H2A and H2B and the
acetylation of their lysines. This raises a question of how the selective acetylation of
histone residues affects the nucleosomes. What are the differences in binding energetics
associated with the site specific residues? Simon et al suggested the possibility of a site
specific response to binding affinity but report only DNA extension as a function of time
as opposed to extension as a function of force, which can provide information on the freeenergy of a rupture event. There is no detailed analysis of binding kinetics for these
histones.
In all of our native and hyper-acetylated histone experiments, I have observed the
sharp peaks at 50nm and 90nm. I have observed a more distinct secondary 90nm peak in
the hyper-acetylated data and attribute that to the inherent instability of these
nucleosomes to forces < 2.5pN. I believe that the instability of these hyper-acetylated
nucleosomes leads to their susceptibility to simultaneous ruptures. I believe that the
cooperative interactions are responsible for the rapid release of two or more nucleosomes
simultaneously. I believe that hyper-acetylated histones are far more reliant on
cooperative interactions. If they are not exposed to forces near f*, they remain stable.
When they approach or exceed f*, one may rupture and the cooperative effect holding the
array together collapses causing a sudden release of all nucleosomes.
The force response histograms show the distinct differences in the binding
affinities of each type of formed nucleosome. I attribute the sharp ~1.6pN peak in figure
4B to be due to the instability of hyper-acetylated histone nucleosome arrays. Although
104
both native and hyper-acetylated nucleosomes peak at the critical force of 1.6pN, as seen
in figure 4A, B, the distribution of native nucleosomes disruption events is spread over a
wider range of force. The sharp peak in 4B is followed by fewer events at higher forces.
This behavior of hyper-acetylated nucleosomes proves that the addition of acetyl groups
greatly reduces the stability of a nucleosome to applied forces above the critical force of
1.6pN predicted by Marko and Siggia. It is important to note that both native and PTM
histones rupture at the predicted theoretical limit.
In a comparison between native, residue-tail-removed, and acetylated histones
undertaken by Brower-Toland et al, a difference in binding affinities was observed, but
no information regarding the binding energetics at low forces <17.6pN was reported
(Brower-Toland et al, 2005). They mentioned only the “smooth energy landscape”
observable at these low forces because the high loading rate intrinsic in their optical
trapping technique prevented them from observing measureable unbinding events. Lowforce, low loading rate data could not be gathered using their optical force clamp or
optical velocity trap. Only a low loading rate in conjunction with a low force observation
allowed us, in comparison, to make these measurements on nucleosomal disruptions.
They present data showing slight shifts in the force response between native, histones
with tails removed, and two classes of histones specified by exposure time to an
acetylating agent. Their data regarding tailless and acetylated histones shows as a shift in
their force-extension curves of a few pN in a force range between 10pN and 30pN. The
slight shift, ranging from 0.6pN to 2.9pN, tabulated in their results is not comparable the
observed step-distribution data I present.
105
4.3.2 Observations on the Results
I present a clear shift in the necessary unbinding forces needed to rupture
nucleosomes formed from native or PTM histones. The data I present shows that the
nucleosome response to forces less than 5pN is far more critical for understanding the
binding characteristics than the 17.6pN to 20pN range that others were studying as well
as forces greater that 10pN where others began to observe the rupturing of individual
nucleosomes.
I have found no other comparable single molecule studies making direct
observations of the differences between native and PTM histones at low forces with
which to compare our results. The experiments mentioned in section 4.3.2 are as close as
I have found.
I present data showing a distinct difference in both the step-sizes and the stepforce relations between native and PTM histones. As I have shown, nearly all PTM
rupture events occur below 5pN. Also, I present data showing that PTM histones are
more susceptible to rupturing simultaneously than the native histone structures. I believe
that this shift is due to the instability of the modified nucleosomes because of the charge
neutralization provided by the hyper-acetylation of the histones. The cooperative
interaction that may exist between the PTM nucleosomes in an array and their
susceptibility to rupturing at very low forces are what I believe are responsible for the
greater prevalence of multiple ruptures. I also suggest the possibility that the homogenous
nature of the hyper-acetylated histones may be responsible for the near complete
106
dissociation of nucleosomes at low forces. The naïve histones can have varying degrees
of naturally occurring PTMs and will have different respective force responses.
I also present data showing that there are differences in the stability of the
nucleosomes. As can been seen in Fig 2B and Fig 5, the unbinding of nucleosomes
formed with PTM histones occur, on average, more rapidly over a smaller window of
force. This rapid dissociation shown in Fig 3B and the force-step histogram of Fig. 4B
suggest an inherent instability of these nucleosomes. I believe that the charge
neutralization provided by hyper-acetylated histones leads to the quantifiable difference
of 0.5s between ruptures and the predominance of the unbinding events occurring at or
below 5pN. With little possible electrostatic attraction left between the histones and the
DNA, they have an increased probability of falling off under any increase in tension. The
native histones show far more stability throughout the force ranges in these experiments,
~1pN through ~20pN, as expected because the charge interactions between the histones
and the DNA is not screened by the addition of nearly as many acetyl groups. Some
PTMs do occur in our native histones, but not enough to neutralize charges as much as
observed in the hyper-acetylated histones.
Chapter 5
Conclusion
5.1 Summary of Results
I have developed a novel magnetic tweezers that advances single molecule
experimentation techniques and incorporates the ability to produce forces of 0.01pN to
~100pN in the plane of focus. This magnetic tweezer is capable of producing near
equilibrium changes in force with an ability to maintain a very low loading rate. With the
simple design of a semi –closed chamber, I can maintain a low-noise environment which
in turn gives us the ability to obtain high-resolution, DNA force and extension
measurements. The simple design also gives me the ability to easily introduce proteins
that bind to DNA.
The in-plane, high-force capability of the optical trap and the simple, fixed-force
characteristics of magnetic tweezers have, until now, remained for almost mutually
exclusive. Optical tweezers were the method of choice to study DNA’s response to a
range of force that covered three to four orders of magnitude and required high resolution
extension data. If control over the applied force and simplicity of design were desired,
then vertical magnetic tweezers would be the most obvious choice.
My goal was to produce an instrument with the ease of use and construction of a
magnetic tweezer and the wide range of force - ~0.01pN to ~100pN - available in optical
traps. I also set out to perform experiments in the plane of focus, like optical traps, in
order to more easily analyze changes in DNA’s end-to-end extension. The horizontal
magnetic
tweezer
that
I
have
developed
107
accomplishes
these
tasks.
108
I then pushed on to use this technique to study the interactions of proteins,
specifically histones, with DNA at various fixed force levels, and have presented data on
nucleosomes, interactions assayed using our method. This technique has allowed me to
successfully detect and control nucleosome binding and unbinding events. With these low
noise and force loading rates, I have presented data that will lead to a better
understanding of the equilibrium binding energetics of DNA-histone interactions.
With this novel, horizontal magnetic tweezers, I have been able to successfully
perform DNA force-extension measurements and observe its DNA force response up to
but not including the overstretching transition, observe the theoretically predicted low
force limit of 1.6pN predicted by Marko and Siggia for the disruption of nucleosomes,
and compare the binding behavior of native and PTM formed nucleosomes showing
~11pN of difference in the binding free-energies.
5.2 Future Directions
The most immediate research goal to tackle is the equilibrium, on-off fluctuation
of nucleosomes below the critical force f * . With the current setup, I have observed what
appears to be equilibrium fluctuations in the formation and disruption of nucleosomes, an
observation not reported previously. With some effort, behavior of nucleosomes
below f * can be observed and analyzed. This new information could give significant
insight into the dynamics of nucleosomes inside the nucleus of cells. In order to make the
109
measurement of these fluctuations one will need to increase the magnification of the
objective used to gather more fluctuation data and increase the frame rate of the camera
from 15frames/s to ~ 150frames/s. The experiment will also need to be altered to provide
a more stable and constant concentration of histones in the solution. Rather than spraying
in a small amount of histones and observing the binding and unbinding events, one will
need to maintain a concentration of histones in solution so as to gather more fluctuation
data at a fixed low level of tension of ~1pN, below the critical f * of 1.6pN. Working
within this force range (<1pN) and using initially with native histone cores, one can also
introduce proteins such as AcetylCoA and Histone AcetylTransferase to investigate the
kinetics and energetics associated with the in-vivo acetylation process, the acetylation
being monitored with extension jumps as histones rupture from the DNA.
Histones in vivo are not typically hyper-acetylated but rather subject to varying
degrees of acetylation. This research project was designed to find the differences in the
stability of nucleosomes formed from native histones and hyper-acetylated histones. I
used histones that had all or nearly all acetylation sites modified with the chemical
addition of an acetyl group. A much more difficult experiment would be one that focused
on the acetylation of specific lysine residues of the histone tails mentioned above. I know
that the binding energy of the acetylated histones is not directly proportional to the
amount of acetylation but rather to the position of the acetyl group on the histone tail.
With the current magnetic tweezer system, one could observe the binding characteristics
of histones that have specific lysines acetylated. The major difficulty of this experiment is
the biochemistry and the biology, rather than the instrumentation and experimental setup.
110
This research would provide a better understanding of the nucleosome and its stability in
conditions that more closely mimic the acetylation of histones in vivo.
An even more ambitious undertaking using the horizontal magnetic tweezer
developed for the experiments in this thesis would be the study of various molecular
motors as DNA polymerase, RNA polymerase etc. and their response to low loading
rates. Our system can make precise measurements of biological nano-machines in action.
Single molecule techniques stand at the forefront of biological research and will
provide invaluable knowledge to our understanding of living systems. These techniques
also push physical and technical barriers in the quest to make better, more precise,
measurements of observable biological systems and structures. Indeed, in the future,
nanotechnology will be needed to advance the understanding of biological systems
I only hope that any future work in this field will be met with the same measure of
collaboration and desire for knowledge that I have been witness to here at The Catholic
University of America.
Appendix I
Tris-EDTA Buffer Preparation 100X
1) Prepare a 150mM Tris solution: Place 18.2mg of Tris in 100mL 0f filtered, sterile
water.
2) Adjust the pH to 8 (use HCl to reduce pH or NaOH to increase pH)
3) Add 0.56g of EDTA to the solution and enough water to make 150mL.
4) Filter the solution using a 0.2µm filter and sterilize in an autoclave
111
Appendix II
DNA End-Functionalization Protocol
DNA comes in buffer solution from New England Biolabs. It is linearized, with 12 base
extensions on both the 3’ and 5’ ends.
Ligating the oligomer 5’ –GGGCGGCGACCT-digoxygenin (ONLdig) to λ-DNA
1. Place 10µg of λ-DNA in a 200µL tube and add ONLdig so that the ratio of oligos
to DNA is 10:1. This ratio can be approximated using the mass of one λ-DNA.
2. Add 1X Tris-EDTA to get a final volume of 35µL
3. Mix and incubate at 65ºC for five minutes to linearize any remnant circular DNA
molecules.
4. Let the DNA cool for five minutes
5. Melt and vortex 10X ligase buffer that is stored at -20ºC
6. Add 4µL ligase buffer to the solution
7. Add 1µL ligase. Try to minimize the amount of time the ligase is out of the -20ºC.
It looses viability rapidly in ambient temperatures.
8. Extract the DNA from the solution using the QIAGEN kit and its protocol.
a. Add 120µL of QX1 buffer and 80µL of water to the 40µL DNA sample.
b. QIAEXII gel needs to be vortexed. Add 20µL QIAEXII to the 220µL
solution.
112
c. Incubate mixture for 10 minutes, gently flicking every minute to assure
proper mixing.
d. Centrifuge for 1 minute. Upon removal from the centrifuge, remove the
supernatant.
e. Wash the sample with 500µL of PE buffer twice.
f. Let the remaining pellet air dry. This can take up to an hour.
g. Add 40µL of Tris-EDTA buffer. This dissolves the pellet.
h. Incubate for 10 minutes at 65ºC
i. Centrifuge for 1 minute.
j. The DNA is now extracted in the supernatant solution. Remove the
supernatant to a clean tube and prepare to attach Biotin oligo.
Ligating the oligomer 5’ –AGGTCGCCGCCC-biotin (ONRbio) to λ-DNA
1. To the 40µL sample of partially functionalizd DNA, add 2µL of a prepared biotin
oligo solution so that there is, again, a 10:1 ratio of oligos to DNA.
2. Mix and incubate at 65ºC for 10 minutes.
3. Let the DNA cool for five minutes
4. Melt and vortex 10X ligase buffer that is stored at -20ºC
5. Add 5µL ligase buffer to the solution
6. Add 2µL ligase. Try to minimize the amount of time the ligase is out of the -20ºC.
7. Incubate for 10minutes at room temperature.
113
8. Extract the DNA from the solution using the QIAGEN kit and its protocol.
a. Add 120µL of QX1 buffer and 80µL of water to the 40µL DNA sample.
b. QIAEXII gel needs to be vortexed. Add 20µL QIAEXII to the 220µL
solution.
c. Incubate mixture for 10 minutes, gently flicking every minute to assure
proper mixing.
d. Centrifuge for 1 minute. Upon removal from the centrifuge, remove the
supernatant.
e. Wash the sample with 500µL of PE buffer twice.
f. Let the remaining pellet air dry. This can take up to an hour.
g. Add 100µL of Tris-EDTA buffer. This dissolves the pellet.
h. Incubate for 10 minutes at 65ºC
i. Centrifuge for 1 minute.
j. The DNA is now extracted in the supernatant solution. Aliquot 5µL into
separate tubes.
k. Store samples in -20ºC freezer until needed for experiments.
114
Appendix III
Labeling of 3µm diameter polystyrene beads with anti-dig-oxygenin
1) Wash 1mL of stock 3-micron polystyrene beads using 1X PBS buffer solution.
2mL of PBS is used in each rinse. The beads are vortexed and centrifuged in the
PBS buffer solution.
2) Re-suspend the beads in a 1X PBS, 5% glutaraldehyde solution overnight in the
refrigerator a 4ºC.
3) Wash the beads at least four times using the method described above.
4) Re-suspend the beads in an anti-dig solution in PBS. The concentration of the
solution is 0.2mg/ml. Incubate for 4 hours at room temperature.
5) Centrifuge the bead-anti-dig solution and discard the supernatant. Re-suspend the
beads in 0.5M ethylolamine in PBS. Incubate for 30 minutes.
6) Centrifuge the ethylolamine-bead solution and discard the supernatant.
7) Re-suspend the beads in a bovine serum albumin (BSA), PBS solution with a
BSA concentration of 10mg/mL. Incubate for 30 minutes at room temperature.
8) Repeat previous step. Store the finished beads in the BSA solution in the
refrigerator for up to 6 months.
115
Appendix IV
Micro-injection protocol.
1) Take a pre-washed glass capillary – see Appendix V for capillary specifications,
storing, and cleaning procedures - and pull a long, straight pipette using the
Narashige PP-83 vertical puller with default settings. The resulting pipette has to
be broken or cracked at a position along its length so that two 10µm – 20 µm
inner diameter tapered pipettes are formed. The location where the pulled pipette
must be broken is determined by trial and error. A small piece of glass capillary or
a glass cutting tile is used to make the break. There is no need to further polish
this pipette using the microforge.
2) An Eppendorf Microloader tip is attached to an Eppendorf pipetter set to draw out
7-10µL of proteins in buffer from an protein solution aliquot. The unused portion
of the aliquot is placed in a 4º C refrigerator. Small amounts, 7-10µL, are used for
a typical experiment.
3) Maintain an air gap on each side of the protein solution in the tip to prevent
dilution of the sample.
4) Place this micropipette in the second, New Era syringe pump system.
5) Set the flow rate to ~2mL/hr.
6) It is very important to push out the air bubble into the buffer-filled sample cell
before the start of the experiment.
116
Appendix V
Protocol for creating and using micropipettes for aspiration.
1) Start with 1mm outer diameter 6 inch WPI glass capillaries.
2) Storing and Cleaning glass capillaries – carry out these steps under a constantly
running hood
a. Glass capillaries are kept fully immersed in a solution of isopropyl,
acetone, and methanol until ready to be used.
b. Before use, take a capillary out of solution and forcefully flush the tube
with isopropyl alcohol. This is done with small diameter rubber tubing and
a syringe.
c. Once flushed, the tubes need to be dried vertically. There is little air
movement inside these tubes; storing them vertically allows any remaining
alcohol the opportunity to drain out.
3) Take a clean glass capillary and place it in the Sutter Instruments P-97
micropipette puller.
4) Using the appropriate program: Heat = 589, Pull = 25, Velocity = 110, Time =
140, two micropipettes with barbed ends are formed. Heat regulates the
temperature with applied current, Pull regulates the strength of the pulling
apparatus, Velocity regulates the pulling mechanism of the instrument to begin as
the glass begins to melt and separate under the applied tension, Ttime regulates
the performance of the cooling mechanism after the pipettes have been pulled.
117
5) Place one of the two new barbed pipettes into the micro-forge.
6) Create a clean, smooth opening with the forge by quickly touching the shaft of the
pipette to a glass bead on the forge’s platinum heating element. As soon as
contact is made between the pipette and the glass bead, turn off the heating
element. This action should, through thermal stress, cause the glass to break
resulting in a correctly-sized opening with a smooth outer surface. If the heating
element is too hot, the pipette will bend away from the element. If the element is
too cool, the pipette may stick, bend, or fail to break.
7) Take the finished pipette to the hood and fill it with 1x Tris-EDTA buffer using a
spinal tap needle. This is done by inserting the needle into the large-diameter open
end of the micropipette and filling. Filling is done by holding firmly the
micropipette while simultaneously applying pressure to the syringe and slowly
withdrawing the spinal tap needle.
8) Still under the hood, take the pipette and connect to the syringe with plastic
tubing. The syringe is filled with 1x Tris-EDTA buffer.
9) Take the syringe, tubing and loaded micropipette, all connected, into the
experiment room and attach to the magnetic tweezer.
a. Place the syringe in the New Era syringe pump
b. Being careful not to break the micropipette tip, place the micropipette in
the clamp that is suspended beneath the Siskyou three-axis micromanipulator.
10) Adjust the Siskyou micromanipulator to find the micropipette above the objective.
118
11) Once found, test to see if there are any blockages in the micropipette through a
visual inspection along the length of the micropipette or by applying pressure to
the syringe. If, when pressure is applied, there is a visible droplet at the tip, it
should work. If performing the test in aqueous buffer, one should see the effects
of fluid flow. Set flow rate or aspiration rate to 5mL/hr or adjust by hand with
syringe pump thumb screw.
12) If any blockage or other hindrance is found, discard tip and start over.
119
Appendix VI
Native Histone Preparation
All proteins were provided by Dr. Tuma of the Department of Biology along with these
protocols.
Reagents and Antibodies
F12 (Coon’s) medium was purchased from Sigma-Aldrich (St. Louis, MO) and fetal
bovine serum (FBS) was from Gemini Bio-Products (Woodland, CA).
Cell Culture
1) WIF-B cells were grown in a humidified 7% CO2 incubator at 37oC as described
2) Briefly, cells were grown in F12 medium, pH 7.0, supplemented with 5% FBS, 10
µM hypoxanthine, 40 nM aminopterin and 1.6 µM thymidine.
3) Cells were seeded at 1.3 x 104 cells/cm2 and grown for 8-12 days until they
reached maximum density and polarity.
Histone purification
1) Confluent monolayers of WIF-B cells grown in 10 cm dishes were lysed in 0.8 ml
of extraction buffer and histones purified using the spin column-based Histone
Purification Mini Kit (Active Motif, Carlsbad, CA) according to the
manufacturer’s instructions.
2) Purity was assessed by analyzing the eluted proteins on Coomassie Blue-stained
SDS-PAGE gels (2).
120
3) The column eluates containing the histones were aliquoted and stored at -80oC.
4) After purification, histone concentrations of ~ 1mg/mL were determined by
densitometric analysis of Coomassie Blue stained gels using bovine serum
albumin as the standard. All of the core histones, H2A, H2B, H3 and H4 were in
solution. The solution did not contain the linker histone H1.
5) The prepared solution was further aliquoted into 5μL samples and stored at
-
20oC.
6) When preparing the samples for an experiment, the small samples would be
diluted to a final concentration of ~ 0.167mg/mL.
The histone core complex was purified from cells using column chromatography.
To assess purity and enrichment of the core complex, the crude histone preparation (CH),
column flow through (FT), wash (W) and the final column eluate (E) were analyzed on
Coomassie Blue-stained SDS-PAGE gels. As shown in Figure 3, the core complex was
efficiently purified from both the cultured WIF-B cells as seen by the robust staining of
the histone subunits in the column eluate. These eluate samples were further diluted in
xx buffer and used in the subsequent DNA binding studies.
121
Appendix VII
Hyper-Acetylated Histone Preparation
All proteins were provided by Dr. Tuma of the Department of Biology along with these
protocols.
Reagents and Antibodies F12 (Coon’s) medium, trichostatin A (TSA) and the HRPconjugated secondary antibodies were purchased from Sigma-Aldrich (St. Louis, MO).
Fetal bovine serum (FBS) was from Gemini Bio-Products (Woodland, CA). The
monoclonal antibodies against histone H3 or acetylated lysine 9 of histone H3 were
purchased from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA).
Cell Culture
1) WIF-B cells were grown in a humidified 7% CO2 incubator at 37oC as described
(1).
2) Briefly, cells were grown in F12 medium, pH 7.0, supplemented with 5% FBS,
10 µM hypoxanthine, 40 nM aminopterin and 1.6 µM thymidine.
3) Cells were seeded at 1.3 x 104 cells/cm2 and grown for 8-12 days until they
reached maximum density and polarity.
122
Western Blotting
1) Cells grown on coverslips were treated with 0, 50, 250 or 500 nM TSA for 30 min
at 37oC.
2) Cells were rinsed 3 times with phosphate buffered saline and lysed directly into
0.5 ml Laemmli sample buffer (2) and boiled for 3 min.
3) Proteins were electrophoretically separated using SDS-PAGE and transferred to
nitrocellulose (2).
4) The membranes were incubated overnight at 4oC with antibodies specific to
histone H3 (1:1000) or acetylated lysine 9 of histone H3 (1:1000).
5) Immunoblots were processed with HRP-conjugated secondary antibodies and
immunoreactivity was detected using enhanced chemiluminescence (PerkinElmer,
Crofton, MD).
Histone purification.
1) Confluent monolayers of WIF-B cells grown in 10 cm dishes were treated in the
absence or presence of 250 nM TSA for 30 min at 37oC.
2) Cells were lysed in 0.8 ml of extraction buffer and histones purified using the spin
column-based Histone Purification Mini Kit (Active Motif, Carlsbad, CA)
according to the manufacturer’s instructions.
3) Purity was assessed by analyzing the eluted proteins on Coomassie Blue-stained
SDS-PAGE gels.
4) Histone concentrations were determined by densitometric analysis of Coomassie
Blue stained gels using bovine serum albumin as the standard.
123
5) The column eluates containing the histones were aliquoted and stored at -80oC.
6) Individual aliquots were thawed immediately before use and further diluted in
Tris-EDTA buffer.
124
Appendix VIII
Derivation of the Fluctuation-Dissipation Theorem
The fluctuation-dissipation theorem used to calculate the magnetic force on the
superparamagnetic particle used in our DNA constructs. In our experiments, a tethered
bead is subject to a unidirectional force, in context to thermal motion. For instance, I
observe a system not unlike a pendulum in our experiments. In a pendulum, I define the
potential energy as a function of applied force and height in the following equation:
U
Fh where h is the height above the potential energy minimum defined as the bottom
of the swing and F is the external force acting on the pendulum, in this example, this
would be gravity. This derivation is courtesy a thesis of Dunja Skoko (Skoko 2006).
The height h is a function of the length of the pendulum L and the angle
associated with the swing
h
according to the following equation:
L L cos
or likewise
h
L(1 cos )
I can redefine the (1 cos ) term with a trigonometric identity
sin 2 ( )
2
1 cos
.
2
Plugging this into the expression for h I get the following:
125
h
2 L sin 2 ( )
2
For small oscillations around the minimum, I can use the small angle approximation
sin
to simplify the last statement so that it can be written as follows:
h
2L( ) 2
2
or likewise
2
h
L(
2
)
Next, the above equation for h can be used in the initial equation for the potential energy
equation U=Fh. With this change of terms, I get the following:
2
U
FL(
2
)
Or likewise,
U
F
(L ) 2
2L
For small angles, L
x so I can write
126
U
F
( x) 2
2L
Above is an expression for the potential energy of small oscillations of a simple
pendulum. This is in essence the same system that is observed in our single molecule
experiments with DNA. To make the leap to a useable expression for our experiments, I
can compare this expression to the average energy of a simple harmonic oscillator which
is given as follows:
k BT
2
U SHO
With the above expression, I can equate the average energy of a simple harmonic
oscillator to the behavior of our tethered bead in the magnetic field by presenting the
following:
k BT
2
U SHO
( x) 2
F
2L
From which we can readily see the final outcome as the following:
k BT
2
F
x
2L
2
And quickly followed by
F
k BT
L
x 2
which is the result I was looking for. In our experiments, the magnetic bead and
the DNA act like the pendulum in this discussion.
127
Appendix IX
Sterilization and cleaning protocols for all Instruments
All bottles and instruments used in the preparation and storing of buffers and all other
pre-experiment protocols need to be cleaned and sterilized to prevent contamination of
DNA, Proteins, beads, etc. The steps in this procedure are as follows.
1) Thoroughly wash with soap and water all necessary instruments and bottles.
2) Thoroughly rinse first with tap water all soap from the bottles and instruments.
3) Thoroughly rinse, three times, with DI water all bottles and instruments to
remove any minerals and chemicals in the tap water.
4) Place all objects in an autoclave, the Harvey Hydroclave Model MC10.
5) Set the program to Unwrapped Instruments and start the process
6) Upon completion, quickly remove items, using thermal gloves, and place in the
hood. The objects are 121ºC when removed.
7) Let cool in the hood. All items are now clean and sterile.
128
Appendix X
Preparation and Construction of Buffer Chamber
All cover-slips and magnets used in the construction of the buffer chamber need to be
cleaned prior to the construction of a buffer chamber and of course an experiment.
1) Take a #1 cover slip and clean, in the sterile hood, with acetone. Spray the coverslip with acetone and wipe with a Kim-wipe tissue.
2) After rinsing the cover-slip with the acetone, spray the glass with isopropyl
alcohol and wipe again wit a Kim-wipe tissue.
3) Take the magnet to be used in the buffer chamber and clean, in the sterile hood,
with acetone.
4) After rinsing the magnet with the acetone, spray the magnet with isopropyl
alcohol and wipe again wit a Kim-wipe tissue.
5) Both are clean and can be used in the construction of the buffer chamber.
Preparation of the Buffer Chamber.
1) Take cleaned cover-slip and magnet and begin assembly of the chamber.
2) Form three walls of the buffer chamber using 2 1cm X 1cm pieces and a 1cm X
3cm piece of cut microscope slides glued to the cover-slip using silicone
adhesive. The arrangement of these three pieces of glass is determined by with the
aid of a simple jig in the lab.
3) The magnet is glued in last. It is placed along the right wall of the buffer chamber
and is fixed in place with minimal silicone adhesive.
129
Appendix XI
DI water filtering and sterilization
All buffers for experiments must be prepared with utmost attention to cleanliness and
sterility. Al buffers are prepared with DI (18 Ohm) water obtained from the second floor
filtration room. After 15L of DI water is gathered, liter bottle samples are sterilized in the
Harvey Hydroclave Model MC10.
1) Fill several one Liter bottles with DI water.
2) Place bottles in the auto-clave.
3) Set program on the autoclave to Liquids and start the sterilization.
4) Upon completion, remove bottles from the autoclave and quiclkly place them in
the sterile hood.
5) Let bottles and water cool in the hood.
6) When needed, filter the sterilized water using a syringe and a 0.2μm syringe
filter, placing the now twice filtered water into another clean, sterile bottle.
7) The DI water is now twice filtered and sterilized, minimizing the risk of
introducing any contamination to the experiments.
130
Appendix XII
Free-Energy Calculation
1) To calculate the average free energy I calculate the free energy released for each
event by multiplying the jump in extension
point f or f i l i
l by the instantaneous force at that
E i . Here, i is the index for the disruption. As an experiment
progresses, starting from 0, we count the number of unbinding events and label
the force at the point of disruption. So, at any jump in extension, the change in
extension and the force are linked together.
2) I then sum the free energy of all individual events.
3) Once all free energies have been summed up, I divide by the total number of
individual nucleosomes associated with the disruption events. This gives us an
average free-energy per rupture;
ni E i
i
i
f i l i . Here, n refers to the
i
number of nucleosomes per event: if
l i is 50nm, n would be 1, if
l i is ~100nm,
n would be 2 and so on. This way, the number of nucleosomes is accounted for as
a measure of change in length,
l.
131
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