I

A Single-Molecule Optical Trapping Assay: Measuring E. coli RecQ Helicase
Motion Using Force-Activated DNA Substrates
byStephen R. Okoniewski
B.A., Northwestern University, 2013
M.S., University of Colorado, 2016

A thesis submitted to the
Faculty of the Graduate School of the
University of Colorado in partial fulfillmentof the requirements for the degree ofDoctor of Philosophy
Department of Physics
2020

II
This thesis entitled:
A Single-Molecule Optical Trapping Assay: Measuring E. coli RecQ Helicase Motion Using
Force-Activated DNA Substrates
written by Stephen R. Okoniewski
has been approved for the Department of Physics________________________________________
Thomas Perkins, Department of Molecular, Cellular, and Developmental Biology________________________________________
Meredith Betterton, Department of Physics________________________________________
Loren Hough, Department of Physics________________________________________
Joel Kralj, Department of Molecular, Cellular, and Developmental Biology________________________________________
Deborah Wuttke, Department of Biochemistry

Date ______________________
The final copy of this thesis has been examined by the signatories, and we find that both thecontent and the form meet acceptable presentation standards of scholarly work in the above
mentioned discipline.

III
Okoniewski, Stephen R. (Ph.D., Physics)
A Single-Molecule Optical Trapping Assay: Measuring E. coli RecQ Helicase Motion
Using Force-Activated DNA Substrates
Thesis directed by Professor Thomas Perkins, Department of Molecular, Cellular, andDevelopmental Biology

In this thesis, I detail the work I have done in studying both force-activated DNA substrates andthe RecQ DNA helicase from E. coli. The first half of the document introduces some backgroundfor these topics as well as the experimental methods used, with a focus on my high-precision
single-molecule optical trap. In the chapter on force-activated DNA substrates, I explain therationale for developing them, prior strides towards completing them, and then our work.
Multiple substrate types are explored and validated, and several single-molecule helicase assays
are done as proof of construct efficacy. The last chapter deals with my research on the singlemolecule unwinding behavior of the RecQ helicase. The main focus of the chapter is optimizing
my assay conditions so I can resolve single base-pair steps of RecQ, in order to better understandits fundamental stepping mechanics and see whether the kinetic models posed in previous papers
are correct. I end the thesis by discussing some novel interactions between RecQ and its DNA
substrate which lead to unexpected unwinding behaviors.

IV

Dedication

I dedicate this thesis to my parents, Robert and Shirliann Okoniewski.
I love you guys.

V

Acknowledgements

I would first like to acknowledge the funding support I received during my graduate career. A
CU/NIH Molecular Biophysics training grant (T32 GM-065103) funded me during my second
and third years, and an NSF Graduate Research Fellowship (DGE 1144083) funded me during
my fourth, fifth, and sixth years. In addition to financial support, these awards also provided
valuable training and professional development opportunities. Thank you for the support you
have given to my growth as a scientist and to the work described herein.
Thank you next to my advisor, Tom Perkins, without whom this thesis would not exist.
His knowledge and experience in the many avenues of scientific life have been immensely
helpful, as have his deep technical and communicative skills.
Thank you to Lyle Uyetake, our group’s research technician, for all the talent and effort
he put into creating the many DNA constructs and helicase proteins I used. Without his help, thiswork would have been so much more difficult.
Thank you to Rebecca Montange, the post-doc who first trained me in using our group’s
optical trapping instruments, wet lab protocols, and code base. Her assistance made my earlydays easier and her wry comments made them entertaining.
Thank you to the post-docs Rob Walder, Devin Edwards, and David Jacobson for serving
as constant sources of advice and humor. And thank you again to Rob for spearheading the azide
functionalization protocol. It improved the reliability of my DNA assays immensely.

VI
Thank you to Matt Siewny and Patrick Heenan, my fellow graduate students in the group,for the advice and understanding they gave while we went through this process together. There
really is nothing like someone going through the same stuff as you to give you solace.
Thank you to Arnulf Taylor, the junior graduate student I have trained to replace me. His
hard work and dedication made the final days of my thesis easier, and I feel confident I’mleaving this project in good hands.
Thank you to the many undergraduates that have aided me in this work. Specifically,
thank you to John van Patten and Ayush Adhikari for the long process of empirically converting
Rob’s azide protocol to my optical trapping coverslips, and for all the times they made azide
coverslips for me. Thank you to Jaeyvn Faulk and Toby Bollig for the time they spent passing on
the knowledge of how to create fiducial mark coverslips.
Thank you to JILA electronics shop wizard Carl Sauer for the many times he built and/or
fixed the electronic components of my optical trapping instrument. He was a lifesaver. Thankyou to JILA Keck Lab manager David Alchenberger for the clean room and SEM training heprovided, and for the times he fixed various components of my instrument.
Thank you to Keir Neuman and Yeonee Seol at the NIH for generously providing mewith RecQ protein aliquots and plasmids, as well as discussions on using and studying RecQ.
And lastly, thank you to Joe Falke and everyone involved in the Biophysics training
program at CU Boulder. As previously stated, the many seminar and presentation opportunities
provided by the program were instrumental to my scientific development, and the biophysics
community that the program fostered has been a wonderful part of my time here.

VII

Contents

Chapter
1

Introduction

1.1

Single-molecule force spectroscopy and optical trapping .................................................. 1

1.2

DNA mechanics and overstretching ................................................................................... 6

1.3

RecQ family helicases....................................................................................................... 13

2

Experimental Methods

2.1

Optical trapping instrument .............................................................................................. 20

2.2

2.1.1

Optical components ........................................................................................... 21

2.1.2

Detection Electronics ......................................................................................... 28

2.1.3

Software communication ................................................................................... 31

2.1.4

Laser stabilization .............................................................................................. 33

2.1.5

Stage stabilization .............................................................................................. 37

Sample chamber manufacture ........................................................................................... 40
2.2.1

Single-channel flow chambers ........................................................................... 40

2.2.2

KOH-etched coverslips ...................................................................................... 43

2.2.3

Fiducial mark fabrication ................................................................................... 43

2.2.4

Azide functionalization ...................................................................................... 46

VIII
2.3

2.4

Biological assays ............................................................................................................... 49
2.3.1

General DNA preparation .................................................................................. 49

2.3.2

Tethered bead assay ........................................................................................... 51

2.3.3

RecQ helicase purification and preparation ....................................................... 52

Data collection and analysis.............................................................................................. 56
2.4.1

Bead position setting and calibration ................................................................. 56

2.4.2

Trap calibration and linearity ............................................................................. 60

2.4.3

DNA tether centering and calibration ................................................................ 65

2.4.4

Overstretching assay .......................................................................................... 67

2.4.5

Active force-clamp assay ................................................................................... 69

3

Force-activated DNA Substrates

3.1

Introduction to force-activation ........................................................................................ 71

3.2

Validation of two force-activated DNA substrates ........................................................... 75
3.2.1

1002 nt 50% GC substrate ................................................................................. 76

3.2.2

Hairpin-unwinding substrate .............................................................................. 81

3.3

Single-molecule measurement of E. coli RecQ on-rate .................................................... 86

3.4

Advanced substrates and %GC-dependent unwinding velocity assay ............................. 89

4

RecQ helicase unwinding

4.1

The physical unwinding step size ..................................................................................... 97
4.1.1

Introduction to RecQ stepping ........................................................................... 98

4.1.2

Optimizing optical trapping conditions ............................................................. 99

IX
4.1.3

Optimizing biochemical conditions ................................................................. 111

4.1.4

Final RecQ stepping results ............................................................................. 122

4.2

Unwinding “popping” behavior ...................................................................................... 130

4.3

Additional non-standard interactions of RecQ and DNA ............................................... 140

Bibliography .............................................................................................................................. 147

Appendix
Acronym / Abbreviation Index ................................................................................................... 155
Protocols Index ........................................................................................................................... 159
Streptavidin to CML beads: Coupling Procedure ........................................................... 160
Cleaning Coverslips ........................................................................................................ 162
Making HSQ Post Coverslips ......................................................................................... 163
Silane-PEG-Azide Functionalization Protocol ............................................................... 165
Tethered Bead Assay: DBCO Edition ............................................................................ 166
RecQ Protein Purification ............................................................................................... 167

X

FiguresFigure1.1

Aspects of optical trapping theory and application..............................................................4

1.2

The basic structure of DNA .................................................................................................8

1.3

DNA stretching and overstretching ...................................................................................11

1.4

The RecQ DNA helicase ....................................................................................................16

2.1

Diagram of the pre-fiber optical setup for each laser ........................................................23

2.2

Diagram of the post-fiber optical setup .............................................................................25

2.3

Labeled pictures of the microscope column and base .......................................................26

2.4

The electronic components of the optical trapping detection system ................................30

2.5

The operation of the AOM servo loop for reducing laser noise ........................................35

2.6

Operation and results of the stage stabilization servo loop................................................39

2.7

Single-channel flow chamber and accessories ...................................................................41

2.8

HSQ-based fiducial mark fabrication ................................................................................45

2.9

Azide functionalization process and accessories ...............................................................48

2.10

RecQ helicase purification gel ...........................................................................................55

2.11

Procedures for positioning and calibrating a bead .............................................................58

2.12

Trap stiffness calibration and linearity ..............................................................................62

2.13

Surface-coupled geometry calculations and DNA centering .............................................66

XI
3.1

Introduction to force-activated DNA substrates ................................................................73

3.2

Forced dissociation of a 1002-nt DNA segment at ~65 pN ...............................................77

3.3

Quantifying force activation in the 50% GC construct ......................................................79

3.4

Challenges and solution to the 20-bp hairpin construct .....................................................82

3.5

Force activation of a DNA hairpin substrate .....................................................................84

3.6

Determining the size of the activated hairpin ....................................................................85

3.7

Force-activation enables an efficient single-molecule on-rate assay .................................88

3.8

Design and application of 120 bp force-activated hairpin substrates ................................93

3.9

66% GC unwinding velocities and validation ...................................................................94

4.1

DNA mechanistic properties and step simulations ..........................................................101

4.2

Surface-coupled geometry optimization ..........................................................................107

4.3

Optimizing the trap stiffness ............................................................................................109

4.4

RecQ processivity varies with [ATP] ..............................................................................112

4.5

RecQ interacts with non-hydrolysable ATP analogs .......................................................115

4.6

Slowing RecQ unwinding with ATPαS and divalent swaps............................................120

4.7

The E. coli RecQ helicase takes 1-bp physical unwinding steps .....................................124

4.8

The high-resolution step size distribution strongly peaks at 1 bp ....................................125

4.9

Fitting simulated magnetic tweezer steps ........................................................................130

4.10

ATP-less RecQ popping of DNA hairpins.......................................................................132

4.11

Quantifying the popping phenomenon.............................................................................135

4.12

Structural implications of the popping behavior ..............................................................137

4.13

RecQ can induce sharp extension drops prior to unwinding ...........................................141

4.14

RecQ can stall force-induced hairpin unzipping ..............................................................144

1

Chapter 1
Introduction

1.1

Single-molecule force spectroscopy and optical trapping

The fundamental unit of life is the cell. From the smallest single-celled bacteria to the largest
flora and fauna, all biological organisms are made up of these basic units. But these units are notsimple. A single cell is an incredibly complex system of hundreds of millions of biochemicalmolecules (e.g., amino acids, nucleic acids, lipids, sugars, both alone and in higher-order
structures) and hundreds of billions of chemical atoms and molecules (e.g., water, ionic salts,
protons), all interacting with each other and the external environment. To understand thefundamental aspects of biology, these physiochemical components and interactions must be
understood. The complexity of this task has given rise to many different scientific fields, eachfocusing on a slightly different facet of life’s basic operation. For instance, many fundamental
biological processes depend upon physical forces and motions between single biomolecules andbiomolecular complexes. This is the purview of my field: single-molecule force spectroscopy.
Single-molecule force spectroscopy (SMFS) is a biophysical field that aims to understandthe dynamics and energetics of biomolecular processes by mechanically manipulating individual
biomolecules. Common topics in the field include investigating the polymer physics of
biopolymers (1-12), the pathways and energy landscapes involved in secondary, tertiary, and2quaternary biopolymer structures (13-23), and the various aspects of molecular motor activity,
including mechanical steps, kinetic rates, dynamic pauses, and stall forces (24-35). SMFS work
is typically carried out using one of three instruments: optical traps, magnetic tweezers, or atomic
force microscopes (AFM). Each has their own advantages, disadvantages, and preferred systems(36), but all three work off of the same idea: attach controllable probes to two ends of abiomolecule, and manipulate those probes to induce and record mechanical forces andextensions. To perform the work described in this thesis, I used an optical trapping instrument.
Broadly, an optical trap refers to any instrument capable of using light-based forces totrap objects in some region of 3D space. Many excellent reviews and books on the theory,
construction, and operation of optical traps exist (37-43). Here I give only a broad outline of thebasic theory, and later describe my particular instrument in detail (section 2.1). The first opticaltraps were pioneered by Arthur Ashkin and coworkers in the 1970s and ‘80s, culminating in thefirst single-beam gradient optical trap in 1986 (44). This type of optical trap (used in my
instrument) is formed by tightly focusing a laser beam to a diffraction-limited spot. This creates afocus with a very high intensity (i.e., high electric field strength, since 𝐼 ∝ 𝐸 2 ) and a steep
electric field gradient surrounding it (electric field strength quickly drops off in all directions). It
is well-established that electric dipole moments are drawn towards regions of high electric fieldstrength in order to minimize the total electric potential energy of the system, so long as themoment and field vectors align (Figure 1.1A, (45) Eq. 4.24). In other words, the electric potential
energy at a point in space can be generally expressed as:
𝑈 = 𝑞𝜙 − 𝒑 ∙ 𝑬 + ⋯ ,(1.1)

where 𝑈 is the electric potential energy, 𝑞 is the net charge, 𝜙 is the electric potential, 𝒑 is theelectric dipole moment, and 𝑬 is the electric field. It is also well-established that an3electromagnetic wave’s oscillating electric field can induce electric dipole moments withindielectric objects, such that the moment and field vectors always perfectly align (so long as thedielectric object’s refractive index is greater than the surrounding medium’s index). This meansthat a tightly-focused laser beam will (a) induce oscillating dipole moments within any nearby
dielectric object, and (b) draw the moments (and therefore the object) towards the beam focus(Figure 1.1B, left). The magnitude of this drawing force depends on the magnitude of theobject’s electric dipole moment and the magnitude of the electric field gradient near the focus.
Because of this, the drawing force is often called the “gradient” force.
Dielectric objects can be trapped within the focal region if this gradient force is greaterthan any other forces on the object, including the scattering force due to radiation pressure fromthe laser beam and the thermal forces due to any external fluid medium (e.g., air or water). The
point in space where an object becomes stably trapped is called the trap center. For a wellaligned laser beam, this trap center is located at the lateral center of the beam (collinear with the
focal point), but just downstream of the focal point axially. This is because the average radiation
scattering force is nonzero (unlike the thermal force), and pushes the dielectric object away fromthe focal point (which would be the true trap center if no scattering force was present) until theaxial gradient force balances the scattering force (Figure 1.1B, right). If the trap is formed byfocusing a collimated TEM00 laser beam with a high numerical aperture (NA) objective lens, theelectric potential energy of the trap will be 3D harmonic near the trap center. This means that,along each axis, the energy and force can be expressed as:
𝑈𝑖 =

1
𝑘 𝑥2
2 𝑖 𝑖

𝐹𝑖 = −𝑘𝑖 𝑥𝑖 ,(1.2)

where i specifies the x, y, or z axis, 𝑥𝑖 is the distance from trap center along that axis, 𝐹𝑖 is theforce, and 𝑘𝑖 is a parameter called the trap stiffness. So long as a dielectric object stays within

4

Figure 1.1: Aspects of optical trapping theory and application.
A) Example 1D plot of electric potential vs position, meant to intuitively demonstrate whyelectric dipoles seeks regions of high electric field. To minimize the electric potential energy ofthe system, positive charges (blue) seek the lowest electric potential, while negative charges(green) seek the highest potential. Electric dipoles (purple) are composed of positive andnegative charges, with the dipole vector pointing from negative to positive. To minimize energy,
the positive part seeks low potential while the negative part seeks high potential. This can be
optimally satisfied by aligning the dipole vector with the largest electric field vector (thenegative gradient of the electric potential). B) Left: A tightly-focused laser beam will induce
electric dipole moments within dielectric objects and draw them towards the high electric fieldof the beam focus (pulling the dielectric object along, as well). Right: For stable trapping, thegradient forces of the trap must be stronger than all other forces, including the scattering force
due to beam propagation. This scattering force will push the object away from the beam focus
until the axial gradient force equals it in magnitude, stably trapping the object and defining thetrap center. C) Basic diagram of a surface-coupled optical trapping assay. Within a liquidmedium, DNA molecules are bound to a glass surface at one end and a dielectric microsphere(bead) at the other. Thermal forces from the medium buffet both the bead and DNA, such thatthe lowest energy state is where the DNA adopts a compact, high entropy conformation (left).
By trapping the bead and moving the stage relative to the trap, the DNA molecule can be
manipulated and stretched. In this geometry, the average force on the bead from the trap is equaland opposite to the force on the bead from the DNA (from both entropic and enthalpic
contributions). By knowing where both the surface and bead are at all times, the force andextension of the DNA can be measured.

5
the harmonic region of the trap (which usually extends at least 50 nm in all directions), its
potential energy and force can be expressed simply through a constant and its distance from thetrap center. Since this region produces a force that is linear with distance, it is also called thelinear region of the trap (outside this region, trap stiffness is no longer constant with distance).
In biophysics applications, the most common object to trap in a single-beam gradient
optical trap is a dielectric microsphere, typically shortened to simply “a bead”. By attaching one
end of a biomolecule of interest to a fixed surface and another end to a bead, the molecule can be
stretched and manipulated by trapping the bead within the optical trap (Figure 1.1C). Spheres areused because they are easy to manufacture and calibrate. This latter trait is especially important,since it is necessary to exert controlled, well-calibrated forces. As an example of the ease of use
enabled by spheres, we can make a rough calculation to determine what variables affect an
optical trap’s linear trap stiffness 𝑘. To begin, from Eq. (1.2) we know that the potential energy
𝑈𝑖 along a specific axis can be parameterized by the trap stiffness 𝑘𝑖 . From Eq. (1.1) we knowthat 𝑈𝑖 at a point in space goes as −(𝒑 ∙ 𝑬)𝑖 . Say the dipole moment is induced in a dielectric
sphere of refractive index 𝑛 immersed in some medium of refractive index 𝑛𝑚 . For ahomogeneous external electric field 𝑬, calculating this moment is a graduate-level exercise:𝑏2 − 1 3
2
𝒑o = 4𝜋𝜖0 𝑛𝑚 ( 2) 𝑟 𝑬,
𝑏 +2(1.3)

where 𝜖0 is the electric permittivity of free space, 𝑏 = 𝑛⁄𝑛𝑚 is the ratio of the sphere and media
refractive indices, and 𝑟 is the sphere radius. The optical trap is certainly not a homogeneous 𝑬
field (if it was, there would be no trapping), but again this is only a rough calculation to obtaindependencies. Note that 𝒑o points in the same direction as 𝑬 only so long as 𝑏 > 1, i.e. so long as
the sphere has a larger refractive index than the surrounding medium (which for most SFMS

6
assays is water-based, so 𝑛𝑚 ≅ 1.33). In addition, we can combine Eq. (1.3) with Eqs. (1.1) and(1.2) to find trap stiffness dependencies:𝑏2 − 1 3 2
𝑏2 − 1 3|𝑈𝑖 | ∝ (𝒑 ∙ 𝑬)𝑖 ∝ ( 2) 𝑟 𝐸𝑖 ∝ ( 2) 𝑟 𝐼𝑖 ∝ 𝑘𝑖 .
𝑏 +2
𝑏 +2

(1.4)

Thus we find three variables that determine 𝑘: the trapped bead’s volume (𝑟 3 ), the polarizabilityof the bead’s dielectric material relative to the medium (𝑏 = 𝑛⁄𝑛𝑚 ), and the intensity of the laser
beam (𝐼). While studies have shown a more complex dependence between bead size and trap
stiffness than the one above (46), up to a point it is true that large beads are better for exerting
large forces. It also turns out that the linear proportionality between trap stiffness and laser
intensity does hold in practice (section 2.4.2). With respect to dielectric material, polystyrene orsilica beads are typically used, as they have 𝑛 values larger than 1.33 and can be functionalizedwith many biomolecule attachment chemistries.

1.2

DNA mechanics and overstretching

In my optical trapping experiments, the tethered biopolymer is almost always DNA, or somemodified form of DNA. Typically, “DNA” refers to double-stranded DNA (dsDNA), which iscomposed of two single-stranded DNA (ssDNA) polymers coiled around each other in a double
helix configuration (Figure 1.2). The monomer unit for ssDNA polymers is the nucleotide,composed of a deoxyribose sugar, a phosphate group, and one of four nucleobases: adenine (A),thymine (T), cytosine (C), or guanine (G). Deoxyribose is a five-carbon sugar, and in a
nucleotide the phosphate group and nucleobase are bound to the 5’ and 1’ sugar carbons,respectively (Figure 1.2, right). Within a single strand, the nucleotide units are joined by covalent
phosphodiester bonds between the 5’ phosphate group of one unit and the 3’ hydroxyl group of
the next. This results in a strong but flexible sugar-phosphate backbone, with all of the7nucleobases pointing outward. Note that a linear, finite strand of ssDNA will have a 5’ phosphategroup exposed at one end and a 3’ hydroxyl group exposed at the other. By convention, we callthe nucleotide with the exposed 5’ phosphate group the first unit in the strand, and the one withthe exposed 3’ sugar the last. We read the base sequence in a strand by reading 5’ to 3’. In a
molecule of dsDNA, the bases between two strands noncovalently bind to each other, with A
bases binding to T bases and C bases binding to G bases. Note that each strand runs reverse to
the other, such that the 5’ unit of one will bind to the 3’ unit of the other. If the strands are
perfect complements (every A/T/C/G base in one binds to a T/A/G/C in the other), then it is only
necessary to specify the sequence of one of the strands in order to know the sequence of both.
The canonical A-T and G-C base pairs, also called Watson-Crick base pairs, areenergetically favorable and stabilize the double helix structure. Part of that stability comes fromthe noncovalent bonds within the base pairs: A-T pairs form via two hydrogen bonds, while G-Cpairs form via three (Figure 1.2, right). Base stacking interactions between the rings of nearest
neighbor bases on the same strand provide a significant amount of additional stability. Fromthese two mechanisms, G-C pairs provide the largest stability, while A-T pairs provide the least(47). On the other hand, non-canonical base pairs (anything not A-T or G-C) are energeticallyunfavorable and disrupt the structure. Much of this disruption is due to geometric constraints inthe double helix. A canonical double helix has a constant diameter of 2 nm (48), since both A-T
and G-C pairs have the same width. Non-canonical base pairs have a larger width, and thus
protrude part of the backbone out from the rest of the helix, straining the structure. If too manyare present (e.g., if the strands of the dsDNA did not anneal correctly), the helix will be too
unstable to survive the constant thermal forces of the surrounding aqueous medium, and will frayapart into its component strands.

8

Figure 1.2: The basic structure of DNA.
Left: Double-stranded DNA is made up of two polymers of nucleotides noncovalently bound ina double helix. A nucleotide is made up of a phosphate group, a five carbon sugar ring, and oneof four nitrogenous nucleobases (Adenine (A), Thymine (T), Guanine (G), Cytosine (C)). Eachsingle strand of DNA is composed of a sequence of these nucleotides, and has a definite polarity(5' to 3'). Center: In a double helix, the strands run anti-parallel to each other, and ideallycontain only Watson-Crick basepairs (A pairs with T and G pairs with C). Right: A moredetailed picture of the chemistry involved in the double helix of DNA. Note that each phosphatein the backbone of DNA has a negative charge, which leads to the full strand being highly
negatively charged. Reproduced from Figure 6 of Lumen's Open Education Resource forMicrobiology: Structure and Function of DNA.
(https://courses.lumenlearning.com/microbiology/chapter/structure-and-function-of-dna/).

9
The polymer properties of dsDNA and ssDNA have been well-characterized and are
critical for interpreting DNA-based data from an optical trap. The most important property is theaverage, zero-tension distance between adjacent monomer units, called the contour length (𝐿c ).
For dsDNA, the contour length specifies the average distance between adjacent base pairs (bp)(also called the helix rise). Early crystallographic studies clearly showed 𝐿dsDNA= 0.338 nm/bpc(49). For ssDNA, the contour length specifies the average distance between the backbone
phosphate groups of the nucleotide (nt) monomer units (since the actual bases can rotate freely).
Single-molecule studies have shown 𝐿ssDNA= 0.56 nm/nt (3, 50). In addition, two other polymercproperties are important for interpreting data: the persistence length (𝐿p ) and the stretch modulus(𝑆). The persistence length characterizes the bending stiffness of a polymer, and is reported as
the average, zero-tension distance at which two units cease to be directionally correlated. For thenegatively charged dsDNA and ssDNA polymers (each phosphate group has −1e charge), 𝐿p
depends on the temperature and ionic conditions surrounding the polymers. At standard salt andtemperature conditions (~150 mM monovalent and 25 °C), 𝐿dsDNA= 50 nm (2, 7) and 𝐿ssDNA=pp0.75 nm (3, 50). The stretch modulus characterizes the enthalpic stiffness of a polymer, i.e. how
difficult it is to forcefully stretch a polymer beyond its zero-tension contour length, and isreported as a characteristic force. At standard conditions, 𝑆 dsDNA = 1200 pN (5, 7) and𝑆 ssDNA = 800 pN (3, 50).
These polymer parameters are used in mathematical models that relate the force (𝐹)exerted on a biopolymer to its resulting extension (𝑥). For dsDNA, the most common model isthe extensible worm-like chain (eWLC). From (5), it is given as:

10

𝐹=

𝑘B 𝑇 𝑥 1 𝐹
1
−
−
+
,
𝐿p 𝐿Tc 4 𝑆
𝑥 𝐹 2
4 (1 − T + 𝑆 )
(
)
𝐿c

(1.5)

where 𝑘B is Boltzmann’s constant, 𝑇 is the temperature, and 𝐿Tc is the total contour length of thepolymer, i.e. 𝐿c multiplied by the number of monomer units in the polymer. Above 𝐹 = 2 pN,
the above expression can be approximated to a simpler one (which I often use) (5):𝑘B 𝑇 𝐹
𝑥 = 𝐿Tc (1 − √+ ).
4𝐿p 𝐹 𝑆

(1.6)

This expression has the benefit of being an explicit function 𝑥(𝐹), as opposed to the implicit
function 𝐹(𝐹, 𝑥) in Eq. (1.5). For ssDNA, an eWLC model or the extensible freely-jointed chain(eFJC) model can be used successfully (51), so long as one does not mix parameters between the
two. I use eFJC models for my data, and from (3), the model is given as:
2𝐿p 𝐹
𝑘B 𝑇
𝐹
𝑥 = 𝐿Tc (coth ()−) (1 + ) .
𝑘B 𝑇
2𝐿p 𝐹
𝑆

(1.7)

These models can fit the force-extension data collected in an optical trap, and thereby interpretits biological meaning. However, it is important to keep in mind that these models do have theirlimitations and quirks. For example, when the force-extension curve from a moderate length
dsDNA or shorter (≤ 1.5 μm total contour length) is fit by the eWLC model, the model will often
report 𝐿p values smaller than expected (11). This is a model quirk, and does not represent any
true change in the polymer’s persistence length.
These polymer models also have limited ranges of applicability. For example, the eWLC
model for dsDNA only works up to ~35 pN (Figure 1.3A), capturing the low-force entropic
stretching (0-10 pN) and the medium force enthalpic stretching (10-35 pN). Specifically, when
the two polymer endpoints are fixed close together (low extension), the polymer can adopt many

11

Figure 1.3: DNA stretching and overstretching.
A) Force-extension curve for dsDNA. When there is no external force applied across a DNA
polymer, it will adopt a tangled, condensed form in solution, maximizing its entropy. It takes
very little applied force to stretch the polymer in this entropic regime, and so the force-extension
curve is very nearly flat. As the extension of the DNA nears its contour length (0.338 nm/bp),
further stretching becomes more difficult, and the necessary force begins to rise dramatically. In
this enthalpic regime, the polymer has no entropic freedom, and further pulling stretcheschemical bonds within the structure. Below ~35 pN, this behavior is well-described by theeWLC polymer model. As the force reaches ~65 pN, the polymer undergoes a dramatic
overstretching event, where the extension rapidly grows by 70%. After this transition finishes, asecond enthalpic regime begins and continues until the dsDNA strands peel apart or a chemicalbond breaks. B) The three mechanisms of DNA overstretching. Red arrows indicate where thetension is being applied. If free ends or nicks are present in the polymer, the strands will begin
to peel apart. This conversion of dsDNA into ssDNA is what causes the extension increase inoverstretching. If no free ends or nicks are present, bases can either internally melt apart or
rearrange themselves into a new, extended form (S-DNA). Both of these mechanisms also
increase the extension to largely the same degree.

12
coiled-up configurations (high entropy). This allows it to easily absorb the buffeting thermal
forces of the aqueous medium, leading to low forces exerted on the fixed endpoints. As the twoends are moved further apart, the polymer can adopt fewer configurations (low entropy), and sothermal forces are transmitted more fully to the endpoints. Correspondingly, more external force
is required to keep the endpoints from moving. Once the tether is pulled nearly straight,
continued pulling stretches the base stacking interactions and backbone chemical bonds of thedsDNA molecule (enthalpic stretching), requiring large increases in force for small increases inextension. The eWLC model predicts that this continues on indefinitely, but dsDNA only follows
this behavior up to ~35 pN. Past this point, the molecule actually begins to soften (moreextension for less force), due to coupling between the actual stretching and helical twisting
induced by the stretching. Applying a correction for this effect to the eWLC enables accurate
modelling up to ~60 pN (52).
In the 60-70 pN range, a dsDNA molecule undergoes an incredible increase in extension,
stretching up to 1.7x its total contour length with little increase in force. This phenomenon, firstobserved in 1996, is called the overstretching transition of dsDNA (3, 4). It took two decades offurther research to reach the consensus that three separate mechanisms are responsible foroverstretching (53-63) (Figure 1.3B): (i) the peeling apart of dsDNA into ssDNA from free ends
or nicks, (ii) the internal melting of base pairs within the dsDNA, and (iii) the conversion of
dsDNA into a new structural configuration called S-DNA (“normal” dsDNA is also called BDNA). Which mechanism(s) take place during the transition depends on the temperature and saltconditions surrounding the dsDNA molecule, as well as its topological structure. For example, atstandard temperature and salt conditions (~25 °C and 150 mM monovalent), a molecule will
undergo the peeling mechanism if any free ends or nicks (backbone breaks) are present (59-61).

13
Near these spots, base pairing and base stacking interactions begin to destabilize and dissolve
due to the high tension, allowing one strand of the double helix to peel away and leave theremaining one under tension. Continual peeling and conversion of dsDNA into ssDNA causes
the dramatic extension increase. If the dsDNA molecule is topologically closed (i.e., does nothave free ends or nicks), peeling is prohibited. With this configuration and high temperatures(~37 °C), the base pair destabilization induced by the force still occurs, but now both ssDNA
strands remain under tension, sharing the load (60, 61). Alternatively, under standard conditions
and closed topology, the force destabilizes base interactions enough for them to shift and reform
into a new, extended S-DNA conformation (the form of which is still debated) (60, 61).

1.3

RecQ family helicases

Within a cell, DNA is continually acted upon by a multitude of proteins. Structural proteinsorganize and de-organize the DNA into higher-level structures like nucleosomes andchromosomes. Replication proteins copy the DNA so that its genetic code can be included inboth copies of a dividing cell. Transcription proteins process the DNA genetic code into RNA
molecules that either get translated into proteins or serve their own roles in the cell. And
critically, repair and recombination proteins remove damage from DNA, allowing all of theproteins above to perform their functions normally. There are many families of repair proteins,
each of which specialize in addressing certain kinds of DNA damage. The protein family ofinterest to this study is the RecQ helicase family.
The RecQ family is a set of proteins conserved from bacteria to humans which unwind
the complex, intermediate DNA structures that form during many processes used to maintain the
genome (64-66). For example, they are known to play a critical role in multiple steps of

14
homologous recombination, a repair process for an often-lethal type of DNA damage called
double-stranded breaks, where nicks in both backbone strands lead to the molecule fissuring into
two separate parts. Due to their important DNA repair role, mutations in human RecQ helicases
are associated with a number of diseases (e.g., Bloom, Werner, and Rothmund-Thomson
syndromes) which, among other symptoms, lead to cancer predisposition and premature aging(64-66). Thus, the RecQ helicase family, one of the major families that make up the SF2 helicase
superfamily (67-69), is both biologically important (malfunction causes disease) and functionallycomplex (unwinding of multiple DNA substrate types). Of the dozen known enzymes in the
RecQ family, the one most of interest to this study is the founding member from Escherichia
coli, simply called RecQ.
Functionally, RecQ is an ATPase and 3’ to 5’ DNA helicase, meaning it hydrolyzes
molecules of ATP for energy and unwinds dsDNA by moving along one of the strands in a 3’ to
5’ direction (70). Early biochemical studies showed that RecQ has ATPase activity on circularand linear ssDNA (meaning that it can translocate along ssDNA, as well) and can unwind bluntend dsDNA at high concentrations (70). It is a mildly processive helicase, in that it can unwind
on the order of 100 bp before dissociating from the polymer (71, 72). A cell mutation study
showed that RecQ activity suppresses illegitimate recombination in E. coli, and that otherrecombination pathways cannot fix the problems caused by removing RecQ (73). A suite ofbiochemical studies found that at 37 °C and low salt concentration, RecQ initiates and disrupts
recombination events (together with RecA and SSB), and forms D-loop and Kappa DNA
intermediates (74); nucleates, unwinds, and catenates covalently closed circular DNA at highconcentrations with the aid of Topo III (75); and displays varying large-scale unwinding activity
depending on ATP, Mg2+, and SSB concentrations (76).

15
The first RecQ crystal structure was published in 2003 using a truncated variant called
RecQΔH (77). It showed that RecQ is a multidomain protein with a catalytic core composed of apair of RecA-like domains, followed by RecQ-family-specific auxiliary domains: a zinc-binding
domain and a winged-helix domain. The final domain of RecQ, the helicase-and-RNaseD–C
terminal (HRDC) domain, had to be removed for the original crystallization but was later
crystallized separately (78) (Figure 1.4A). The two RecA-like domains are common to the SF1and SF2 helicase superfamilies, and are used for ATP hydrolysis and ssDNA translocation (67,68). A high degree of conformational flexibility has been observed between the two domains,
specifically during different steps of the ATP hydrolysis cycle (79) (Figure 1.4B). The domains
are observed to rotate relative to one another along an axis defined by the ATP binding cleft; it isthought that this rotation is how the helicase steps along ssDNA, transducing ATP energy into
“inchworm” translocation. The mechanics of this translocation for 3’ to 5’ SF1 helicases are
known: the HD2 domain remains tightly bound to ssDNA during the ATP binding step of the
kinetic pathway. This causes the ssDNA to slide across the HD1 domain during the subsequentrotation caused by ATP binding, and puts the domain in a position to bind a downstream
nucleotide. After the hydrolysis step of the pathway, it does exactly that, tightly binding thenearest nucleotide while HD2 domain releases its previous nucleotide and binds a nearer one.
Through this process inchworm steps are taken (79). Later studies expanded on these structures
to show that in RecQ an aromatic-rich loop couples ATPase and translocation activities in the
catalytic core (80) and that the winged-helix domain serves as a binding pocket for SSB (81, 82),
in addition to modifying core activity.
Recently, crystal structures of RecQ family proteins bound to DNA substrates have beenobtained, a daunting task for the last two decades (79, 83, 84). While E. coli RecQ has still not16

Figure 1.4: The RecQ DNA helicase.
A) The structure of RecQΔH (PDB 1OYW) and its HRDC domain (PDB 1WUD). Full RecQ has
never been crystallized. The two RecA-like motor core domains are dark and light green, thezinc-binding domain is red, and the winged-helix domain is blue. The zinc ion appears as a grey
sphere with the zinc-binding domain. B) RecQ translocates along ssDNA in the 3' to 5' direction
through an ATP hydrolysis cycle. Two motor core structures are shown: E. coli RecQ withATPγS bound (grey; PDB 1OYY) and human RECQ1 with ADP bound (purple; PDB 2V1X).
Figure reproduced from Vindigni et al., 2010 with permission from Elsevier. These structures
show how the left domain (D1) of the motor core changes between the ATP and ADP phases of
the hydrolysis cycle, relative to the D2 domain. This motion drives RecQ translocation. C) The
crystal structure of C. Sakazakii RecQ bound to a forked DNA substrate. The winged-helix
domain undergoes a conformational change to bind the dsDNA portion of the substrate, while
the ssDNA portion appears to be pulled away at a 90° angle (PDB 4TMU).

17
been crystallized with a substrate, another bacterial RecQ from C. sakazakii has been crystallizedwith a forked ss-dsDNA substrate (83). The catalytic core of this protein (CsRecQΔH) has ~86%
sequence homogeneity to EcRecQΔH, making it the closest homolog to E. coli RecQ in the RecQ
family. Interestingly, the protein appears to bind its substrate by rotating its winged-helix domain
and keeping the DNA fork bent at ~90° (Figure 1.4C). Unlike many other helicases, including
some RecQ family proteins, there is no beta-sheet hairpin or other structural motif present to aid
unwinding. Thus, dsDNA unwinding seems to occur only due to the tension placed across the
double helix as the RecA-like domains pull it out of the binding pocket via its attached ssDNA.
The mechanistic implications of this structure are still being explored.
One topic of interest for E. coli RecQ is whether it binds and unwinds DNA as amonomer (single protein) or multimer (many connected RecQ proteins working in concert). The
first paper on the topic found that at 0 mM monovalent and 1 mM divalent salt, high temperature(37 °C), and high concentration (100 nM), RecQ helicase activity is cooperative with ATP
concentration (Hill coefficient 3.3), indicating a multimer of at least three helicases (76).
However, a different group found that between temperatures of 4–37 °C and up to concentrationsof 20 μM, RecQ exists as a monomer both in solution and during unwinding (85-87). A latercollaboration between this group and another lab found RecQ monomers have a “functional”
cooperativity at 37 °C and modest-to-no cooperativity at 25 °C; this functional cooperativity can
result in Hill coefficients greater than 1 despite the sole presence of RecQ monomers (88).
However, the first group revisited the subject, found a Hill coefficient of 2.1 for ssDNA
translocation activity, and then posited that translocation requires at least a dimer (89) (with nodiscussion of the result of (88)). In a recent paper, they also assert that unwinding involves a hostof multimeric species (90). Thus, the multimeric status of RecQ is still an active area of research.

18
Another topic of interest is the fundamental physical, chemical, and kinetic step sizes of
RecQ during ssDNA translocation and dsDNA unwinding. The physical step size of a helicase isdefined as the physical distance it travels during a single reproducible cycle of motion. For
translocation, this means the number of nt it can stably step across, and for unwinding this means
the number of bp it unwinds. The chemical and kinetic step sizes are defined similarly, butinstead of being per cycle-of-motion, they are explicitly per ATP hydrolyzed and per ratelimiting step of activity, respectively. Whether or not these three values are the same or different
depends on the helicase. The first bulk measurement of the unwinding chemical step size of
RecQ found a value of 1 bp (87) (corrected from their original result of 4 bp). Others bulk
studies found that both the translocation and unwinding chemical step sizes are very close to 1
(71, 72). However, there have been observations of a 5 bp kinetic step size for RecQ, the detailed
mechanics of which are still unknown (91). In addition, a recent single-molecule FRET assay
gave the first measurement of RecQ’s unwinding physical step size. Contrary to expectations,this assay appeared to show RecQ taking steps of multiple sizes, from 0.5 bp to 3.5 bp (92).
Thus, finding the true unwinding step size of RecQ is still an active area of research.
The dynamic stepping behavior of RecQ is also an active area of research, and includes
phenomena of interest such as the pauses taken during unwinding, the potential for backsliding
on DNA substrates, the prevalence of strand switching during unwinding, and the effects that
substrate geometry, RecQ mutations, and buffer conditions have on the above. For instance, arecent magnetic tweezers study showed that the HRDC domain induces significant pausing on
hairpin substrates, but only minor pausing on gapped, linear substrates (while not affecting the
instantaneous unwinding rate at all) (93). Strand switching, where a single RecQ moves along
one strand of a dsDNA substrate only to abruptly transfer over to the other strand, is posited to

19
be an important mechanism for how RecQ rescues stalled replication forks (94). It is expectedthat precise single-molecule methods like optical traps will be able to further elucidate these
dynamic phenomena, which are difficult to observe using classical ensemble assays.
Especially important to this work are the similarities and differences between native
RecQ and the truncated protein RecQΔH. As stated above, the HRDC domain induces pauses ofvarying intensities, meaning that the truncated RecQΔH experiences fewer pauses and thus has ahigher mean unwinding rate than its native structure. RecQΔH also has ~2x higher ATPase
activity and ~3x lower ssDNA binding strength than native RecQ (72, 93). On the other hand,the ssDNA binding on-rate is roughly the same between the two (95), as well as the couplingstoichiometry, processivity, and instantaneous unwinding rate (72, 93).

20

Chapter 2
Experimental techniques

In this chapter, I detail the experimental techniques I used in my thesis work. First, I go throughthe basics of my surface-coupled optical trapping instrument, including the active stabilization
methods used to give it sub-nm precision (section 2.1). Next, I cover the fundamental part in allof my samples, single-channel flow chambers, including the various ways we treat and modify
coverslip surfaces (section 2.2). This is followed by the biological work that goes into readying
biophysical experiments, including DNA preparation and tethered bead assays (section 2.3).
Finally, with the instrument and samples explained, I detail the process of performing anexperiment, including the alignment and calibrations needed as well as my basic measurement
assays (section 2.4).

2.1

Optical trapping instrument

In this section, I give a basic overview of my optical trapping instrument. I first go through theoptical components that make up the majority of the physical instrument, including the lasers, thefree-standing optical elements, and the inverted microscope in which experiments are performed(section 2.1.1). I next detail the detection electronics, including the quadrant photodiodes(QPDs), the JILA electronic boards, and the National Instruments (NI) data acquisition

21
electronics that communicate with the instrument’s computer (section 2.1.2). I then cover the NI
software used to collect data and control the various parts of the instrument (section 2.1.3).
Finally, I explain in detail the operation of the servo loops used for laser stabilization (section
2.1.4) and stage stabilization (section 2.1.5).
Note that all of the instrument components are located in an acoustically quiet basement
room (NC30) that is temperature regulated (±0.2 °C) with its own separate airflow. The
temperature is typically set to 22 °C. All of the free-standing optical components are installedinside plexiglass boxes on floating optics tables to insulate them against environmental noise. Allof the electronic boards are mounted in and powered by a NIM rack.

2.1.1 Optical components
The first components pertinent to the instrument are the lasers we use for trapping and detection.
The trapping laser beam is generated by a solid-state neodymium-doped yttrium orthovanadate(Nd:YVO4) laser (Spectra-Physics, Millennia IR, 𝜆 = 1064 nm, 10 W). For high power andlasing stability, the laser is set to operate at 7.8 W output power (7.77 W set on the laser
controller). For additional intensity stability, the cooling system in this laser was modified: theinternal fan was replaced by an 18.6 L/s external fan (connected via custom-built air ducts) that
operates continuously instead of in cycles. The laser emits a single-mode, Gaussian TEM00
beam; this mode is ideal for trapping, as it can be focused to the smallest diameter beam waist,and therefore produces the strongest, most harmonic trap. Two diode lasers are used to produce
the detection laser beams for the instrument. A 𝜆 = 808 nm laser (Lumics, 100 mW) makes thebeam that monitors the position of optically trapped objects, and a 𝜆 = 845 nm laser (Lumics,
200 mW) makes the beam that monitors the position of fiducial marks on the coverslip surface of

22the sample chamber. Lab tradition refers to these lasers as the 810 nm and 850 nm, respectively.
Both diode lasers are operated via custom-built electronic temperature/current controller boxes,
which are primarily used to set the diode current in each laser and stabilize the diode’s
temperature. The beams of both lasers are sent through fiber Bragg gratings to increase theirwavelength stability and optically isolate them from back reflections, such that they both emit asingle-mode TEM00 beam. The wavelengths of these three lasers were chosen for theircommercial availability, relative transparency in water, biological safety, and band separation.
This last characteristic is critical, given that all three beams have to be combined and later
separated using dichroic mirrors.
Since lasers often emit beams which are not fully linearly polarized, the polarization ofeach beam is purified directly after emission using a half-wave plate and a polarizing beam
splitter (PBS) cube. PBS cubes reflect one direction of linear polarization and transmit the other,so we use a half-wave plate to shift the polarization of the laser output so that maximum power is
put into our direction of choice (Figure 2.1; transmitted for the diode lasers, reflected for thetrapping laser). This fully polarized beam is then sent into an acousto-optic modulator (AOM),the fundamental component of the active stabilization loop that reduces laser noise and sets the
final laser intensity in the sample plane. The servo loop will be covered in more detail in section
2.1.4. For now, it is enough to know that the AOM acts as a tunable diffraction grating that
outputs a 1st-order beam of variable power. This beam is coupled into the rest of the system
while the 0th-order and higher order beams are discarded. Each laser beam is then coupled into apolarization-maintaining fiber-optic patch cable (OZ Optics) using a converging lens and a 5axis ultra-stable fiber coupler (Thorlabs PAF-X-5-B). This cable, another part of the AOM servo
loop, propagates the beam to the main part of the optical setup.

23

Figure 2.1: Diagram of the pre-fiber optical setup for each laser.
A) The 810 nm bead detection laser setup. B) The 850 nm post detection laser setup. C) The
1064 nm trapping laser setup. Note the 850 nm and 1064 nm beams are resized using two-lenstelescopes. Additionally, all beams are focused onto their fiber launches to mitigate pointing
noise, especially the 1064 nm beam. This figure was taken and modified from Figure 2.1 ofAmanda E. Carpenter's thesis from the Perkins lab. Acronyms: AOM (acousto-optic modulator),
PBS (polarizing beam splitter).

24
Each beam is launched out of its fiber-optic patch cable using a free-space fiber launchsystem (Thorlabs KT110). A converging lens and PBS cube are rail-mounted to this system; thelens recollimates the beam and the PBS cube repolarizes it. Each beam is then passed through a
90/10 beam sampler, so that 10% of its power is sent into an analog PIN photodiode detector for
the AOM servo loop (Figure 2.2). The 1064 nm and 810 nm beams are then passed throughelectronic shutters which can quickly block or unblock the beams. Likewise, the 1064 nm and
850 nm beams are reflected off of electronically controllable PZT mirrors (PI S330.2SL PZTmounts) so that their positions within the sample plane can be precisely adjusted (the 810 nm
beam is kept fixed in the sample plane, and is used as a point of reference). All three beams arethen combined together using dichroics, passed through a two-lens telescope to resize their beamwidths, and sent into an inverted microscope (Nikon TE2000-S). Entry into the microscope
forms a sort of dividing line within the instrument. Everything before it, including the as-yet
described AOM servo loops (section 2.1.4), all serve to produce three laser beams with minimal
intensity, mode, position, and polarization noise, whose intensity and position can be preciselychanged by the experimenter. Those three beams are the sole outputs of this section of the
instrument, and they now act as inputs into the microscope section.
A diagram of the critical microscope components is given in Figure 2.2, while a labeled
picture of the microscope is given in Figure 2.3. The microscope’s base and condenser column
are reinforced with custom-built monolithic aluminum slabs to improve their mechanicalstability. Additionally, custom-built monolithic parts are used as reinforced mounts for theobjective lens and sample stage. The objective lens is an oil immersion, high numerical aperture(NA) lens (Nikon CFI Plan Apochromat 100x oil IR, 1.4 NA, working distance 0.13), as is thecondenser lens above the stage (1.4 Nikon HNA Oil). The sample stage is a 3-axis PZT stage (PI25

Figure 2.2: Diagram of the post-fiber optical setup.
This diagram shows the beam paths of the 1064 nm (red), 810 nm (orange), and 850 nm (green)
beams. This figure was taken and modified from Figure 2.2 of Amanda E. Carpenter's thesis
from the Perkins lab. Acronyms: CCD (charge-coupled device), ND filter (neutral density
filter), PZT (lead-zirconate titanate piezo-electric transducer).

26

Figure 2.3: Labeled pictures of the microscope column and base.
A) Right side of the microscope and part of the plexiglass box that contains the post-fiber, freestanding optics. B) Left side of the microscope and part of the closed path leading to the CCD
camera. Acronyms: Al (Aluminum), CCD (charge-coupled device), PZT (lead zirconate titanate
piezo-electric transducer), QPD (quadrant photodiode).

27
P-517.3CD) with a microscope slide mount attached to its top, and is controlled by a digitalpiezo stage controller (PI E-710.P3D). At the top of the microscope column, a liquid light guideconnects to a mercury arc lamp located several feet away. With appropriate filters in place, thislamp shines 532 nm light down the column, through a dichroic mirror in the detection arm thatreflects infrared light and passes visible light, and into the sample plane. The lamp is kept far
from the microscope to prevent its heat from inducing mechanical noise. Beneath the objective
lens, another dichroic mirror reflects the three infrared laser beams into the objective lens and to
the sample plane. It also passes the visible 532 nm light from the sample plane to a CCD camera(DAGE-MTI CCD100S) that is attached to the side of the microscope. This camera records the
sample plane image and sends it to a monitor screen to be displayed.
As stated above, the laser beams couple into the microscope column using a dichroic,
which reflects them towards a high NA objective lens. The two-lens telescope they pass throughto enter the microscope shapes the 1064 nm beam width so that it just overfills the back apertureof the objective (~8 mm diameter). This ensures the 1064 nm beam is focused tightly in thesample plane, and thus achieves its highest trapping strength. In addition, the objective lens andthe two PZT mirrors (for the 1064 nm and 850 nm beams) are spaced such that the plane definedby the objective back aperture is optically conjugate to the planes defined by the mirror surfaces.
This means that pure rotations in one plane (e.g., the PZT mirror plane) produce pure translationsin the other (e.g., the aperture plane). This propagates to the sample plane, so that pure rotationsof the PZT mirrors result in pure translations of the 1064 nm and 850 nm beams in the sample
plane; this is critical for accurate beam placement and later calibrations.
The beams are focused by the objective lens, propagate to the sample plane where they
achieve their tightest focus (and where experimental samples are mounted and manipulated), and28
continue on into the condenser lens. During experiments, the free space between the objective
lens and the sample, as well as the sample and the condenser lens, is filled by immersion oil(Nikon type A oil, n = 1.515) to achieve better beam propagation. Also, the objective andcondenser lenses are aligned for Kohler illumination, which provides the sample plane with themost isotropic illumination from the mercury arc lamp. While propagating though the sample
chamber, the beams will partially scatter off of dielectric objects. The scattered portions of thebeams propagate onward with the unscattered portions, and they interfere with each other (96).
This interference is critical for object detection, as will be discussed below. The beams emerge
collimated from the condenser and reflect off of the dichroic mirror into the detection arm of the
microscope column (Figure 2.4A). This detection arm is an optical cage system that includes two
dichroic mirrors and three quadrant photodiodes (QPDs). The two dichroics separate the three
beams into three different beam paths (undoing the combination performed before entering the
microscope). A QPD detector sits at the end of each of these beam paths. The operation of thesedetectors, and the electronic manipulations that are performed on their signals, will be covered in
the next section, as this is the point where the optics part of the instrument ends and theelectronics one begins.

2.1.2 Detection Electronics
The quadrant photodiode (QPD) at the end of each beam path has a planar photon detection area
divided into four equal quadrants. The photon intensity incident on each quadrant is converted
into a separate analog voltage signal and output through stress-relieved cables. The QPDs
themselves are spaced along the beam paths (z position) so that back-focal plane detection ispossible, and their perpendicular (xy) positions are set so that the non-scattered, incident beam

29hits each quadrant with equal intensity. Three-dimensional position detection of microscopic
dielectric objects is possible due to the interference between scattered and unscattered light whensuch an object partially blocks the beam’s propagation path (96). If the object is positioned offcenter from the beam axis in xy, the scattered and unscattered light will interfere such that thebeam intensity distribution is shifted in xy. These shifts are detected by the QPD, as the intensity
on each quadrant changes (while the total incident intensity remains the same). On the other
hand, objects that move along the beam path (z position) shift the overall intensity (96, 97), but
not the xy intensity distribution. The calibrations necessary to convert these intensity changes
into real-space position changes are described later (Section 2.4.1).
Each QPD’s four voltage signals are then sent to a separate custom-built JILA circuitboard that performs a number of signal manipulations. This includes turning the quadrant signals
into x, y, and z signals and passing them through a low-pass anti-aliasing filter. If we name thequadrants counterclockwise from the top right 1, 2, 3, and 4 (Figure 2.4B), then the z signal isjust the sum signal (V1 + V2 + V3 + V4) while x and y are (V1 + V4 – V2 – V3) and (V1 + V2 – V3 –V4), respectively. The x and y signals are normalized by the sum signal (z signal) to ensure they
stay independent of sum signal fluctuations. The low-pass filter frequency is set to half the dataacquisition frequency to prevent aliasing when the signals are later converted from analog todigital. The board then outputs the xyz signals (Figure 2.4C).
The x and y signals go straight into a BNC breakout box that passes the signals into a
connector block (NI SCB-68) and to our PXI data acquisition (DAQ) and control system. Thissystem consists of a PXIe chassis/controller (NI PXIe-1082) that powers and buses data between
a number of electronic DAQ boards. The x and y signals are sent into a multifunction inputoutput (IO) board (NI PXIe-6368) that samples and converts them from analog to digital using a

30

Figure 2.4: The electronic components of the optical trapping detection system.
A) Top view of the detection arm containing the three QPD detectors. B) Face-on view of a
QPD, with the four detection quadrants labeled. C) Custom-made JILA electronic boards
installed into a NIM rack. Above is one of the BNC breakout boxes used to condense signals
into a single, multi-channel NI cable. Acronyms: AOM (acousto-optic modulator), BNC(Bayonet Neill–Concelman connector), NI (National Instruments), NIM (nuclear
instrumentation module), QPD (quadrant photodiode).

3116-bit analog-to-digital converter (ADC). It receives its sampling frequency from another board
in the chassis, the PC communication board (NI PXIe-PCIe8375), that transmits commands fromthe instrument’s computer to the chassis. This communication process will be discussed more in
Section 2.1.3. The 16-bit ADC digitizes the full range of the signal channel, which is 20 V (-10to 10 V) on this instrument, and assigns a 16-bit integer to each sampled voltage. This gives asampling resolution of 0.3 mV. For good spatial resolution, we want to make sure that thevoltage change which arises from objects moving ~1 Å is much larger than 0.3 mV. Happily, forthe xy signals it is 7x–8x larger.
However, this is not true for the z signal, as the sum signal voltage fluctuations due to z
motion are extremely small. Thus, the signal must undergo an addition stage of processing before
it can be sent to the multifunction IO board. From the summation and filtering board, the z signal
is sent to another custom-built JILA electronics board which offsets and amplifies the signal. The
offset is used to zero the signal so it can maximally amplified, and the amplification gain is
usually set to 1000x. After amplification, the z signal is sent to the same breakout box as the xy
signals, and follows the same path to the ADC board. Once sampled and converted, the now
digital signals are bused to the PC communication board and sent into the PC.

2.1.3 Software communication
All electronic communication between the PC and the rest of the instrument is done through thecoding platform LabVIEW (National Instruments). In LabVIEW, one can create a wide variety
of programs (called VIs, or Virtual Instruments) for real-time control and data processing. As
seen in the last section, all measured analog voltages are eventually sent to the PXIe chassis to bedigitized and directed to the computer. LabVIEW is the platform that both requests collection of

32
these signals and serves as their final destination. Data transfer between LabVIEW and the PXIe
chassis is mediated by another NI program, the Measurement & Automation Explorer (MAX).
This program serves as a device manager, and is the link between the NI DAQ hardware (thechassis and its various boards) and the NI software (LabVIEW). Thus, data collection follows
this general workflow: A LabVIEW VI is written to specify which voltage channels one wishes
to read and at what sampling frequency. During an experiment, this VI runs on the computer andsends the task to MAX, which sends the task to the PXIe chassis, which sends the task to themultifunction IO board. The IO board samples the analog input voltages and digitizes them, thensends them back through the same pipeline until they are collected and recorded in LabVIEW.
There they can be processed in real-time, used to make control decisions for other parts of the
instrument, and saved to disk.
In addition to recording input signals, LabVIEW also serves as the output signal source
for controlling various parts of the instrument. The same basic workflow applies: code is writtenthat requests an output voltage signal be sent. When run, the code communicates the task throughMAX to the chassis and multifunction IO board, which then converts the digital signal to ananalog output voltage using a digital-to-analog converter (DAC). The signal is sent to a port on a
BNC breakout box, which connects to part of the instrument. This is how the 1064 nm and 810
nm shutters are opened and closed, how the 1064 nm and 850 nm PZT mirror mounts are rotated(via the PZT mount controller), and how the intensity control on the 1064 nm AOM servo board
is adjusted (to give real-time trap stiffness control).
LabVIEW is also used to control the PZT stage. However, this type of communication
does not involve MAX as an intermediate. Instead, we use low-level VIs originally created by
the stage’s manufacturer to send commands from LabVIEW to the stage controller. This type of

33
communication uses the GPIB protocol (General Purpose Interface Bus) along a USB cablestraight from the computer. GPIB is well-established and reliable, but uses ASCII strings to send
information. These strings limit signal speed, which can be problematic in control applications
such as active stabilization. The resolution to this issue is addressed in section 2.1.5. However,for all other purposes GPIB communication is fast enough, with the benefit of also being simple.

2.1.4 Laser stabilization
The laser beams we use for trapping and detection, while stable by manufacturer standards, are
not inherently stable enough to differentiate between noise and the small signals arising from Åscale biophysical motions. Therefore, we use stabilization servo loops to actively remove noisefrom each laser system. Laser beams can suffer from a variety of noise sources: intensity noise,polarization noise, mode noise (i.e., power in non-TEM00 modes), and pointing (i.e., beam
position) noise. The stabilization servo loop is based on a simple idea: convert as much noise as
possible into intensity noise, and then use an acousto-optic modulator (AOM) to mitigate that
intensity noise. Below I detail the operation of this feedback loop and its various components.
Note that while each of the three lasers in this instrument has its own loop with its own settings,
in operation they are almost exactly the same.
The first and last component of the servo loop is the AOM (Figure 2.5A,B). It consists ofa crystal attached to a piezoelectric transducer. When an AC signal is applied to the transducer, it
rapidly expands and contracts at the driving frequency. These vibrations hit the attached crystal,producing a travelling sound wave within it (which is made up of sections of more and less dense
matter). Such sections possess higher and lower refractive index, respectively, and so act like adiffraction grating to any light passing through the crystal. The amount of light diffracted from34
the incident beam (0th order) into higher order beams (1st order and larger) depends on theamplitude of the sound wave, which is set by the amplitude of the AC voltage signal driving thetransducer. This signal is produced by a separate device, the AOM’s modulation driver, and has afixed frequency of 80 MHz (changing the driving frequency moves the diffracted beams inspace, which is not something we want). This is how the AOM acts as a tunable diffraction
grating that outputs a 1st-order beam of variable power (as previously stated in section 2.1.1).
This 1st-order beam, which can have a power between 0 and 75% of the incident beam, iscoupled into the rest of the system while the other beams are discarded. Thus the intensity of the
final beam can be rapidly changed by changing the amplitude of the driving voltage. This is how
the servo loop will compensate for intensity noise (Figure 2.5C gives the full diagram).
The next servo loop component is the polarization-maintaining fiber-optic patch cablethat couples the 1st-order beam. We use an angle-cleaved input connector to reduce reflections,and a flat output connector to ensure the beam emerges in a Gaussian TEM00 mode. The diodebeams are focused towards their cables using a single converging lens, while the 1064 nm setup
uses two lenses (Figure 2.1). These lenses mitigate the pointing noise that arises during large
servo changes. To understand how this noise arises, note that using a large drive voltage induces
thermal heating in the AOM crystal, expanding it. This expansion changes the spacing of thediffraction grating (akin to changing the driving frequency). This moves the 1st-order beam,hence pointing noise. While the fiber cable propagates the beam to the main part of the opticalsetup, its most important job is converting pre-fiber pointing and mode noise into post-fiber
intensity noise. This occurs because both noise sources reduce the input coupling efficiency,
meaning less light gets into the fiber. Specifically, pointing noise moves the beam away from theoptimal fiber coupling spot, and mode noise puts some beam intensity into non-TEM00 modes

35

Figure 2.5: The operation of the AOM servo loop for reducing laser noise.
A) Side-view cartoon of the AOM. The RF drive signal drives the PZT transducer (black box) tovibrate, which propagates into the crystal (dashed lines) as a sound wave (dotted lines). The
active aperture (dark grey region where the sound wave and material aperture overlap) candiffract any beams (red) that pass through it. B) Top-view cartoon of the AOM. Maximum
diffraction occurs when the input beam enters the AOM at the Bragg angle. C) Diagram of
AOM servo loop beam path and components. D) Top-view picture of the initial components ofthe servo loop. The half-wave plate and PBS cube (beam block out of shot) send a linearlypolarized beam to the AOM, which is mounted onto a 4-axis tilt aligner for easy Bragg anglealignment. Reproduced from Okoniewski et al., 2017 with permission from Springer Nature.
Acronyms: AOM (acousto-optic modulator), PBS (polarizing beam splitter), PZT (leadzirconate titanate piezo-electric transducer), RF (radio frequency).

36
which do not propagate well. At fiber output, the beam is recollimated and passes through a PBS
cube. This cube turns any polarization noise into intensity noise, since only light intensity withthe correct polarization passes through.
With mode, pointing, and polarization noise converted into intensity noise, part of thebeam is siphoned off by a 90/10 beam splitter and sent to a PIN analog photodiode detector(YAG-444AH). These photodiodes are set up to have 60 MHz bandwidths and 5 ns rise times for1064 nm incident light when a large –180 V reverse bias is applied (reverse biasing silicon
photodiodes helps eliminate wavelength-dependent filtering effects, which are especiallyprevalent at 1064 nm). We reduce this reverse bias to –30 V, which gives the photodiodes more
thermal stability at the cost of decreased detection bandwidth and response times. However, wefind this bias to be a good compromise since the servo loop bandwidth is already limited by theAOM, as discussed below. The photodiode converts the incident beam intensity into a current,
which is transformed into a voltage by an attached trans-impedance amplifier.
The photodiode signal is sent to a proportional-integral servo circuit board (Figure 2.4C).
This board works to keep the photodiode voltage (the real laser intensity) equal to a predetermined reference voltage (the requested laser intensity). It does this by modulating its only
output, the voltage it sends to the AOM modulation driver, whenever the photodiode voltagechanges. Changing the input voltage to the AOM modulation driver changes the driving signal
amplitude it sends to the AOM transducer, which thereby changes the intensity of the diffracted
1st-order beam. This is the mechanism by which laser stabilization is achieved: all beam noise isconverted into intensity noise and compensated for by changes to the AOM diffraction
efficiency. The reference voltage used by the servo board determines the final beam intensity inthe sample plane; for the diode lasers, that voltage is internally set in the board, since detection

37
beam intensity does not need to be changed during an experiment. For the trapping laser, thatvoltage is determined in LabVIEW by the user and sent out through the pathway described in
section 2.1.3. This design enables rapid changes to the trap stiffness during experiments.
The bandwidth of this servo loop is ultimately limited by the time it takes to propagatechanges in driving signal amplitude across the AOM crystal via sound waves. This limit will
depend on the beam diameter size inside of the crystal (Figure 2.5), since more time is needed topropagate the sound waves across larger beams. We use beam diameters ~2 mm in size, which
gives us a servo bandwidth of ~200 kHz.

2.1.5 Stage stabilization
In a surface-coupled optical trapping instrument, the system under study (usually DNA withmodifications or associated proteins) gets attached to the sample chamber surface at one end andan optically trapped bead at the other. Position drift of the surface relative to the trapping anddetection beams stretches or relaxes the DNA tether. These length changes convolve into andmask real biophysical signals, reducing the overall signal-to-noise ratio (S/N). Therefore, in
addition to stabilizing laser beam noise, we often add an additional level of stabilization for highresolution assays: active stabilization of the sample surface with a 3D PZT stage.
The idea behind stage stabilization is simple: we add rigid, dielectric fiducial marks to thesurface of our sample chambers so that their position can be read by the 850 nm detection beam.
If the fiducial mark (and therefore the surface) drifts away from a set position within the beam, aservo loop calculation is made to move the surface back into position using the PZT stage. The
construction of these post-like fiducial marks is detailed in section 2.2.3. This section focuses on
the operation of the servo loop itself.

38
To stabilize the surface during an experiment, the following general steps are performed(Figure 2.6A). Right before data is about to be taken, the 850 nm detection beam is roughly
centered over a fiducial mark using its PZT mirror. A LabVIEW VI is then used to precisely
center the beam over the mark and calibrate its voltage response into real-space positions. Oncethe beam is centered and calibrated onto a mark, another VI records the mark’s position inside
the beam at one instant of time. The VI then continually takes the difference between that
position and the mark’s current position. If that difference is nonzero, a correction signal sent to
the PZT stage. The simple version of the servo loop uses proportional correction, where theposition difference is multiplied by a constant gain factor to produce the correction signal. Moreadvanced servo calculations, like integral or differential corrections, are not used. This original
version also uses GPIB communication, the ASCII-based protocol described in section 2.1.3 thatissues all normal PZT stage commands. This servo loop calculates and sends the proportionalcorrection within a LabVIEW VI, and runs at a typical bandwidth of 100 Hz. With this servo
loop, the stage position can be stabilized with sub-nm precision in all three axes (Figure 2.6B).
While this proportional, software- and GPIB-based servo loop is sufficient for many
applications, we also developed a more advanced version, modified from previous work (98).
The major improvement is that servo calculation and stage command operations are moved fromcomputer-based software to circuit board hardware. Specifically, the servo loop calculations are
run on an FPGA (Field Programmable Gate Array) circuit board (NI PXI 7854R) installed within
our NI chassis. An FPGA is a circuit board whose electronic components can be programmed
into a variety of circuit diagrams. This programmability makes the FPGA a versatile tool for
performing extremely fast calculations. A computer is needed to initially program the board, butotherwise the FPGA runs independently using its own internal clock, enabling much faster and39

Figure 2.6: Operation and results of the stage stabilization servo loop.
A) Diagram of the beam path and components of the stage servo loop, including the choice ofusing either a software-based (computer) or hardware-based (FPGA) servo calculation method.
Reproduced from Okoniewski et al., 2017 with permission from Springer Nature. B) 100 Hz 2
beam 1 post data. All gains 0.1. Note that z axis separately amplified. C) Allan deviations ofstabilized parts of traces in (B). Acronyms: FPGA (field programmable gate array).

40
more deterministic operations than computer-based software. The FPGA also enables PIO(parallel input-output) communication with the stage controller, as opposed to GPIB. The PIO
protocol uses simple binary signals for commands instead of GPIB’s large ASCII characters,
enabling faster communication. Another improvement is that an integral correction term is now
calculated in addition to the proportional one; while proportional calculations are reactionary(i.e., they respond to immediate drift), integral calculations try to preemptively prevent drift byobserving drift trends (from past proportional responses) and correcting for them in advance. Allof these improvements combine to make a servo loop that runs deterministically at 500 Hz (nearthe resonance frequency of the stage, the current bandwidth limit) without software-based
interruptions or timing limitations.

2.2

Sample chamber manufacture

In this section, I discuss the sample chambers I use in all of my experiments. I first cover theirgeneral design and construction (section 2.2.1), then go through the various ways we modify thecoverslip surface. First, I cover the protocol for cleaning and etching coverslips (section 2.2.2).
Next, I detail the process of fabricating fiducial marks onto the surface for high-resolution assays(section 2.2.3). Finally, I go through the protocol for azide functionalizing coverslips so that they
can be used in click-chemistry reactions (section 2.2.4).

2.2.1 Single-channel flow chambers
The single-channel flow chambers used in my experiments have a simple design and can beassembled at a lab bench by hand in a few minutes. An example chamber is pictured in the
middle of Figure 2.7. The single channel has dimensions of 25 x 5 x 0.15 mm3, which

41

Figure 2.7: Single-channel flow chamber and accessories.
Left: A custom-machined Teflon microscope slide mount. The two black dots near the top
indicate where the tape edges should be placed. Center: An example single-channel flowchamber, constructed out of a microscope slide, a coverslip, double-sided sticky tape, andepoxy. Right: A Teflon coverslip holder with detachable handle. Reproduced from Okoniewski
et al., 2017 with permission from Springer Nature.

42
corresponds to a volume of ~15 𝜇L. The channel is used to flow various buffers and reagents intoand out of the chamber. The method used to create surface-bound DNA tethers in these flowchambers is given in section 2.3.2. I typically build anywhere from two to six chambers on an
experiment day, depending on the amount of data I want to collect.
To build a chamber, we first insert a microscope slide (Corning 75 x 25 x 5 mm3) into acustom-machined Teflon mount (pictured on the left side of Figure 2.7). We next cut a ~7 cm
long piece of double-sided sticky tape (Scotch, ½ inch wide) in half lengthwise. Using tick marks
drawn onto the Teflon mount as a guide, we stick the two tape halves to the slide 5 mm apart,
perpendicular to the slide’s long axis. To ensure smooth sticking, we press the tape down into theslide with a pair of rubber-tipped tweezers. We then take a cleaned/modified coverslip (Corning
22 x 40 mm2, thickness 1 ½) and lay it along the two tape halves, again orthogonal to the slide’s
long axis. To ensure smooth sticking and remove air pockets, we remove the slide from themount and use the tweezers to press the coverslip into all parts of the tape. We then use arazorblade to cut off any excess tape overhanging the coverslip. These steps are repeated asneeded to make the number of desired sample chambers. We then mix together 5-minute epoxy(Devcon) to rigidify the chambers, applying it to all gaps between the slide and coverslip (except
for the two channel openings). The liquid epoxy wicks into the gaps between the surfaces andhardens, providing a rigid, liquid-tight, single-channel sample chamber.
The single channel is composed of a coverslip surface bottom, microscope slide top, andsticky tape sides. Unlike the coverslip, the microscope slide is not cleaned before use, nor is any
treatment performed on the sticky tape. And while no contamination has ever been observed
from these components, it is common practice to rinse the channels of newly-constructedchambers by flowing through 200 𝜇L buffer (typically 20 mM NaCl, 20 mM HEPES pH 7.5). In

43
addition, note that the channel dimensions are extremely large compared to the tethered beads
commonly trapped in the instrument. This means that the coverslip surface is the only surface tohave a hydrodynamic effect on the solution surrounding the beads (see section 2.4.1).

2.2.2 KOH-etched coverslips
Before coverslips can be modified by fiducial mark fabrication or azide functionalization, theymust first be cleaned and etched. Cleaning removes contamination and etching smooths thesurface to enable more consistent, isotropic functionalization reactions and improved nonspecific protein binding (e.g., antidigoxigenin; see section 2.2.4). Note that etching must be
performed before fiducial mark fabrication, since the etch can partially dissolve the marks.
We use a concentrated solution of 3 M KOH (potassium hydroxide) dissolved in 70%
ethanol and 30% water for microscopic cleaning and etching. A beaker of this solution in placedin a sonicator, as well as a beaker of acetone and two beakers of 18 MΩ purified water. The
sonicator is used to enhance the diffusion and impact of the solutions’ molecules. The protocol
has the coverslips (placed in the Teflon holder shown to the right in Figure 2.7) submerged
sequentially in the beakers of acetone, KOH, water, water for 3 min intervals. After eachinterval, the coverslips and holder are washed down with either an ethanol or water squirt bottle.
After the four beakers, the coverslips are squirted down once more with water then ethanol, driedin a microwave, and stored in parafilm-wrapped containers.

2.2.3 Fiducial mark fabrication
For the best stabilization performance, we want symmetric fiducial marks that are rigid andstrongly attached to the coverslip. This way, detected drift is due to surface drift and not relative

44
motion between the mark and surface. To this end, we manufacture fiducial marks out ofhydrogen silsesquioxane (HSQ; Dow Corning FOx-16), a negative resist solution that crosslinks
when exposed to concentrated electron or extreme ultraviolet (EUV) beams. By coating acoverslip with HSQ and selectively irradiating cylindrical sections of the coating with acontrolled electron beam, we can induce only those sections to crosslink. Once that is done, thecoverslip can be submerged in a TMAH (tetramethylammonium hydroxide) solution to etchaway the noncovalent HSQ while preserving the covalent sections. These cylindrical sections are
the fiducial marks, and due to their shape we commonly refer to them as “posts”.
The fabrication process outlined above can be divided into three parts: coat the coverslip
with HSQ, crosslink HSQ sections with an electron beam, and remove the unwritten HSQ(Figure 2.8). We coat coverslips using a spin coater, a small mount which can spin rapidly andhold anything placed onto it using vacuum suction. We place a coverslip onto it, start the spin
process (2000 rpm for 45 s), and immediately pipette 200 𝜇L HSQ into the middle of thecoverslip. The spinning isotropically spreads the liquid HSQ over the entire surface, which
ensures that the fiducial marks all have the same final height (typically ~600 nm for 200 𝜇L
HSQ). We then place each coverslip onto a 180 °C hotplate (dry side down) to harden the HSQ
by partially dissolving its volatile carrier solvent. This makes later manipulation and transporteasier and ensures the coating keeps its even thickness.
To crosslink HSQ sections, we beam-write with a scanning electron microscope (SEM).
This instrument use an electron beam gun to image and write, and is equipped with a lowvacuum sample chamber that uses water vapor to de-charge dielectric samples. Charging occurs
when a sample cannot conduct away the bombarding electrons from the SEM, i.e. non-metallic
samples. Excess sample charge can deflect the incoming electron beam and generate many

45

Figure 2.8: HSQ-based fiducial mark fabrication.
A) The general steps for fabricating glass-like fiducial marks onto a glass coverslip: clean thesurfaces, spin on the HSQ, irradiate the HSQ with an electron beam to create the posts, anddevelop away the excess HSQ. B) An example image of a fabricated coverslip with posts, taken
by AFM. Reproduced from Okoniewski et al, 2017 with permission from Springer Nature.
Acronyms: AFM (atomic force microscopy), HSQ (hydrogen silsesquioxane), SEM (scanningelectron microscope), TMAH (tetramethylammonium hydroxide).

46
undesirable effects, and is a serious concern in SEM work. Previous versions of this protocol did
not have access to a low-vacuum SEM to automatically de-charge the HSQ, and so used analuminum coating on top of the HSQ to de-charge the sample. Happily, that is no longer
necessary, and we simply insert an HSQ coverslip into the SEM mount after some preliminaryalignment steps. After further time aligning SEM components, we run a program (written in thedevelopment platforms DesignCAD and NPGS) that beam writes a 1.5 x 1.5 mm2 square array ofcylindrical posts into the HSQ, ~600 nm in diameter spaced 15 𝜇m apart. Once done, thecoverslip is removed from the SEM chamber and the process is repeated as necessary.
The development process is very simple. All beam-written coverslips are loaded into a
Teflon coverslip holder (the same kind used in the KOH treatment) and submerged in a TMAH
development solution (Dow electronic chemicals MF CD-26, 2.4% TMAH) for 15 min. This islong enough to remove all of the non-crosslinked HSQ from the coverslips, leaving an array ofcylindrical posts. The coverslips are then washed with water and dried with nitrogen gas.

2.2.4 Azide functionalization
Surface-coupled experiments require one end of a DNA molecule to be attached to the sample
chamber coverslip surface. In early experiments, this was accomplished through a noncovalentdigoxigenin / antidigoxigenin interaction. Specifically, a solution of 100 𝜇g/mL antidigoxigenin
antibodies in PBS 1X was flowed into the single channel and incubated for 1 h. During this time,
the antibodies nonspecifically bind to the coverslip surface, coating it with potential digoxigeninbinding sites. After a buffer wash, 75 𝜇g/mL of bovine serum albumin (BSA) was then incubatedin the chamber for 1 h to nonspecifically bind to and passivate the remaining patches of glass
surface. After another wash, 300 pM DNA with a digoxigenin tag on one 5’ end was incubated47
for 2 h; the dig binds to the antidig, coupling the DNA to the surface. While this method is astandard way to attach DNA to surfaces, it has many clear problems: it relies on nonspecificsticking, noncovalent bonds, and multiple incubations.
Azide functionalization addresses all of these problems (99). In this method, silane-PEGazide molecules (Nanocs Inc. PG2-AZSL-600 or PG2-AZSL-3400, where the end number
specifies the molecular weight) are covalently bound to the surface, making it both nonstick (viathe PEG) and available for covalent bioconjugation. When DNA molecules with DBCO
(dibenzocyclooctyl) labels attached to one of their 5’ ends are incubated with these surfaces, the
DBCO and azide groups undergo a copper-free click chemistry reaction to produce a covalent
linkage. The DNA is therefore attached to the surface via two covalent linkages and a PEG
molecule of selectable size. The azide functionalization process is detailed below (Figure 2.9).
Once functionalized, these coverslips are stored dry at 4 °C in a parafilm-sealed container. They
retain their full efficiency for up to two weeks, and a workable efficiency for up to a month.
To begin the functionalization process, cleaned coverslips are UV irradiated to improve
the reaction efficiency between the silane (SiH4) and the coverslip surface (SiO2) groups. This isdone by incubating the coverslips in a UV ozone chamber (Novascan Technologies, PSD-UV8)
for 30 min. Next, the coverslips are submerged in a ~0.15 mg/mL solution of silane-PEG-azide
dissolved in toluene. For this step, we machined a custom Teflon mount that held the coverslips
and displaced a large volume of fluid (Figure 2.9). With this mount, up to 12 coverslips can befunctionalized at a time using only 100 mL of the silane-PEG-azide solution. The coverslips areleft submerged in this solution in a covered beaker for 30 min at 60 °C (fluid temperature) on a
hot plate. This incubation time and temperature were empirically determined to give the best
functionalization results. Once the incubation is done, the coverslips are sequentially washed48

Figure 2.9: Azide functionalization process and accessories.
A) A custom-machined Teflon coverslip holder for the azide functionalization process. The
holder was designed to displace a large amount of fluid, so the amount of silane-PEG-azide per
coverslip used is minimized and overall costs reduced. A coverslip is placed in the second-leftslot as an example. B) The steps of the functionalization process. A circular AFM coverslip is
depicted as an example; in practice, the above Teflon holder is used with rectangular coverslips.
Adapted from Walder et al., 2017 with permission from the American Chemical Society.
Acronyms: PEG (polyethylene glycol), UV (ultraviolet).

49
with toluene, isopropanol, and 18 MΩ purified water. The initial transfer between the silanePEG-azide solution and the toluene is performed rapidly to prevent drying (which can create
ruinous PEG agglomeration). The coverslips are then removed from the block holder, transferred
to a regular holder, and dried with nitrogen gas.

2.3

Biological assays

In this section, I discuss the biological preparations that go into surface-coupled optical trapping
experiments. I first detail some of the general steps that go into preparing the micron-scale DNA
used in my experiments (section 2.3.1). I next cover how to prepare the fundamental assay for
my trap, the tethered bead assay (section 2.3.2). I then go through the storage and preparation
procedures for the E. coli RecQ helicase proteins studied in this work (section 2.3.3).

2.3.1 General DNA preparation
In my experiments, I tend to work with DNA molecules 3000–6000 bp long, as they aregeometrically easy to use in my instrument. And while much of my work utilizes speciallyengineered DNA regions, these regions never exceed ~1000 base pairs (bp) in length. The
remaining length of each molecule, the relatively inactive “handle” regions, are derived from an
M13mp18 plasmid (New England Biolabs). A complete DNA molecule is typically composed of
two handle regions and one engineered region; often, the engineered region is contained within
the handle primers. These regions are combined using matching endonuclease sites and ligation,
after each individual region has been amplified by polymerase chain reaction (PCR). The basic
workflow for building a complete DNA molecule is this: PCR amplify all component regions,cut them with their endonucleases, and then sequentially ligate pairs of the products together.

50
Then the full product is inserted into a plasmid, and biotin- and DBCO- labelled primers are used
to amplify the construct. The following paragraphs give a detailed workflow.
The first step to building a complete molecule is to PCR amplify its component regions.
We perform the PCR using a Bio-rad T100 Thermal Cycler, and typically use either KOD hot
start polymerase (Novagen 71086-4) or Taq polymerase (Life Technologies 10342-053). DNA
primers are bought from Integrated DNA Technologies. While the primer sequences and PCR
conditions vary from construct to construct, all PCR products are purified using a Qiaquik
purification kit (Qiagen 28106), then loaded onto a 1% agarose/TAE (Tris-HCl, Acetic acid,
EDTA) gel where the product band is excised. The product is then processed by a Bio-Rad
Freeze ‘N Squeeze (Bio-Rad 732-6165), concentrated with a Millipore Amicon Ultra 0.5 mL
10K filter column (Millipore UFC501096) with TE (Tris-HCl, EDTA) buffer exchanges, thensubjected to another Qiaquik kit to remove residual agarose.
The purified PCR products are then cut with their respective endonucleases, exposing
complementary ssDNA “sticky ends” that can anneal to one another. Two of the products are
combined together in equimolar ratio so that they anneal, then are ligated with T4 DNA ligase(New England Biolabs). Ligations are incubated at 16 °C for 16 h in 50–100 𝜇L volumes, with
5% ligase (the maximum amount possible) and 1–2 pM of each DNA. The ligated product isthen run on a 1% agarose/TAE gel and the band is excised and purified as above. If a third PCR
product exists, the process is carried out again between that component and the previous ligation
product. Fully ligated constructs are then incubated with Taq polymerase at 72 °C for 10 min to
generate A overhangs for TA cloning.
To make PCR amplification of this final construct easier, it is cloned into a bacterial
plasmid using TOPO® XL PCR cloning kit (Life Science K4700-10). The plasmid is transformed

51
into competent E. coli, grown up in Luria Broth, and then isolated (Qiaprep Spin Miniprep Kit,
Qiagen 27106). After verification by sequencing and PCR, these plasmids are used as templates
for the biotin- and DBCO-labeled primers. The steps for that PCR are the same as above.

2.3.2 Tethered bead assay
To perform experiments in a surface-coupled optical trap, DNA must be attached to the sample
chamber surface at one end and a dielectric bead at the other. This section describes the protocol
used to generate those attachments, specifically a covalent azide-DBCO bond from the DNA to
the surface and a noncovalent streptavidin-biotin bond from the DNA to a bead. The assay
requires sample chambers (section 2.2.1) made with azide-functionalized coverslips (section
2.2.4), as well as streptavidin-coated beads. I prepare such beads using a well-establishedprotocol (100) which covalently coats polystyrene beads (r = 250 or 375 nm, Invitrogen /Molecular Probes, C37241) with streptavidin (SA) molecules (Vector Laboratories, SA-5000).
To begin, the DNA stock must be diluted in a DNA deposition buffer and incubated in
the sample chamber. However, we have found that the exact DNA and salt concentrations
needed to produce good samples varies depending on the batch of silane-PEG-azide in use. We
assume this batch variation is due to varying levels of trace chemical contaminants produced
during silane-PEG-azide synthesis affecting the electrostatics near the sample surface. Roughly,
we have found that DNA concentration values of 0.1 nM to 1 nM work, with monovalent salts of
20 mM to 100 mM. Our current protocol uses these conditions: 1 nM DNA in a buffer of 20 mM
NaCl, 20 mM HEPES pH 7.5, and 1 mM EDTA pH 8. At times, I have found it helpful to add 2
mM MgCl2 and 2 mM EDTA pH 8, as well, but this varies batch-to-batch. We prepare 15 𝜇L ofthis dilute DNA solution per chamber. Once made, this solution is set aside and 200 𝜇L of

52
dilution buffer (without DNA) is flowed through each chamber using aspirator suction to clean
the flow channel. The DNA solution is then flowed in and the samples are left to incubate inhumidity chambers for 1 h at room temperature.
During this incubation, a 2.5 pM solution of SA-coated beads is prepared from 250 pM
stock. To remove any free streptavidin present in this bead solution (which diffuses much fasterthan the beads and could bind to the DNA first), a buffer exchange is performed. 1 𝜇L of SAbead stock is mixed with 100 𝜇L of a buffer we call MTWB-150 [20 mM Tris-HCl (pH 7.5), 150
mM NaCl, 0.4% Tween-20], spun down in a centrifuge at 14 krpm for 5 min, and its supernatantremoved. The beads are then reconstituted in 100 𝜇L MTWB-150 and inserted into a sonicatorfor ~40 min to ensure monodispersity (clumped beads attached to a tether cannot be used). The
assay is timed so that bead sonication ends at the same time as the DNA incubation. The sample
channels are then washed with 200 𝜇L PBS 1X and 200 𝜇L MTWB-150 to remove the nonbound DNA, and 15 𝜇L of the SA-bead solution is flowed into each sample. The sample are thenincubated for 20 min at room temperature so that the SA-beads can diffuse and bind to the biotinlabels on the surface-bound DNA molecules. After this incubation, the chambers are washedwith 200 𝜇L MTWB-150 and 200 𝜇L working buffer (whatever that happens to be for theexperiment at hand) to remove the excess beads.

2.3.3 RecQ helicase purification and preparation
Initially, we did not make the RecQ proteins ourselves, but instead received stock aliquots fromYeonee Seol and Keir Neuman at the National Institutes of Health. The method they used tocreate these proteins has been detailed (101). Specifically, we were gifted 8.5 µM protein
aliquots containing 30 mM Tris-HCl (pH 7.5), 200 mM NaCl, 1 mM BME, and 10% glycerol,

53
and stored them at –80 °C. The first batch we received worked very well; however, I found that
subsequent RecQ batches had much lower unwinding activity, indicating that part of the batch
contained inactive / misfolded proteins. After trial and error checks between ourselves and ourcollaborators, we still could not identify the exact reason for the lower activity. We then decided
to make the RecQ proteins ourselves, with Yeonee and Keir kindly providing us the plasmids tomake wild-type RecQ and the HRDC-deleted RecQ truncation. Specifically, our lab technician
Lyle did the wet work in producing the proteins, while I researched and compared different
RecQ production methods and tested the batches for activity.
Lyle followed the protocol in (101) exactly when making our first batch of RecQΔH, andthe final product turned out exactly like the recent aliquots we had received from the NIH: lower
activity than expected. My standard method of testing RecQ activity is a single-molecule opticaltrapping assay where I check whether the frequency of unwinding at 200 pM RecQ (at saturatingATP) is comparable to our initial gold-standard aliquots and prior on-rate measurements (further
detailed in section 3.3). However, to ensure these new RecQ batches were tested accurately, I
also performed a bulk ATPase assay with the help of the Leinwand lab at CU Boulder. That
assay confirmed the results of my single-molecule activity assay: the recent RecQΔH batchesfrom the NIH and our newly created one were performing poorly.
Lyle and I worked to debug the problem. Comparing the purification protocols ofdifferent groups showed a lot of small variations in buffers and storage conditions, but those did
not improve our end product. I then noted that RecQ has a zinc-binding domain, which meansthat each RecQ protein needs one ion of zinc to fold properly. Perhaps the low observed activity
was due to a resource shortage: there being many more recombinant RecQ proteins in the E. coli
cells than there were available zinc ions. If that was the case, many of these expressed proteins

54
would not fold properly, and so lose their activity. To address this, Lyle first switched the cell
growth media from Luria broth to Terrific broth, which has 5x more yeast extract in it (and thuszinc). This batch had more activity, though it is hard to tell whether the additional nutrients were
truly the deciding factor. I then found a RecQ protocol that expressed the proteins at 18 °C for 16
h instead of at 37 °C for 4 h (102). Cool protein expression is generally done to ensure correct
folding, and since misfolding was our hypothesized problem source, we immediately made abatch with Terrific broth and 22 °C (room temperature) expression. Cool protein expression
should also result in fewer proteins being expressed per unit time, which would help alleviate
zinc scarcity in folding if that turned out to be the problem. This batch’s activity turned out very
nicely, comparable to our original gold standard batch, and so we kept and adapted this protocol(provided in the Appendix). Figure 2.10 gives an example SDS-PAGE gel of both purified
RecQWT and RecQΔH.
To prepare an unwinding experiment, I first prepare a tethered bead slide (section 2.3.2).
During the final bead incubation step, I remove a protein aliquot from storage, thaw it on ice, anddilute it down using a dilution buffer [30 mM Tris-HCl (pH 7.5), 100 mM NaCl, 10% glycerol].
Aliquots of ATP and DTT, stored at –20 °C, are thawed simultaneously. ATP is needed to fuelthe RecQ protein’s processive translocation and unwinding activity; DTT provides a reducing
environment that can aid in protein stability (though I have found RecQ performs equivalently
without DTT in solution). These components are used to make a working buffer containing 30
mM Tris-HCl (pH 7.5), 100 mM NaCl, 1 mM MgCl2, 0.4% Tween-20, 1 mM DTT, and thedesired final ATP and RecQ concentrations.

55

Figure 2.10: RecQ helicase purification gel.
An SDS-PAGE gel of RecQΔH (lanes 1 and 2) and RecQWT (lane 4). Lane 3 is the size standard.
Lane 1 is the initial 6xHis-TEV-RecQΔH (obtained after step II.7 in our RecQ purification
protocol in Appendix B). Lane 2 is the purified RecQΔH after the tag has been cleaved off with
TEV protease and a negative selection performed. Lane 4 is corresponding result for RecQWT.
This gel assay was performed and recorded by Lyle Uyetake, the Perkins lab technician.

56
2.4

Data collection and analysis

In this section, I discuss the basic protocols we use to calibrate signals and perform experimentsin the optical trap. First, I detail how we position a trapped bead (either free or attached to a
DNA tether) in space and calibrate its detector beam voltage signal to its real space position(section 2.4.1). I then cover the methods we use to determine the optical trap’s stiffness 𝑘T and
its linear region (section 2.4.2). Knowing both parameters is necessary to accurately measure theoptical force exerted on a trapped bead. These first two sections thus explain how we measure abead’s position and force, our two fundamental measurements. Next, I go through how we center
the surface-DNA attachment point directly under the trapped bead and simultaneously determine
the rough worm-like chain (WLC) parameters for the DNA tether (section 2.4.3). I finish bydiscussing the two main types of experiments run on this instrument: overstretching assays(section 2.4.4) and force-clamp assays (section 2.4.5).

2.4.1 Bead position setting and calibration
To set the position of a trapped bead, one first has to actually trap a bead. This is fairly easy forbeads attached to DNA tethers, as the surface can be imaged using the CCD camera. To do this,we use the fact that the trapping and detection beams reflect a small portion of their intensity offof the coverslip glass as they transmit through the chamber. This reflection hits the CCD camera
and is strongest when the beam focus is on the surface. From this surface position, tethered beads
can be seen and it is simply a matter of moving the stage so that a bead gets pulled into the trap.
Free beads, on the other hand, can diffuse to any location in the sample chamber. To trap one ofthem, we move the stage (and thus sample chamber) down so that the trap focus goes deep into

57
the chamber (we find ~6 𝜇m above the surface is a productive location). We then wait for a beadto diffuse by and move the stage to trap it.
With a bead trapped, its average position will now match the position of the trap center.
For the best detector response, we want this trap center to coincide with the center of the 810 nm
detection beam. We thus need to center the lasers so that they are collinear. To do this, we use
the 1064 nm beam’s PZT mirror to move the beam in xy space (the 810 nm beam stays fixed).
Recall that rotating the PZT mirror translates the beam in the sample plane. The mirror can berotated either by hand (for large translations) or by the PZT mount controller, which we directfrom the computer. Using a standard calibration procedure, sub-nm calibration precision can beachieved for controller-directed mirror motion (the procedure is detailed in A. E. Carpenter’s
thesis, section 2.4.8; it involves using a fiducial mark and the factory-determined, sub-nm
position precision of the PZT stage).
If the trapping and detection beams are far from each other, we first roughly center thetrapping beam over the detection beam by hand (using the reflections on the CCD camera); thistypically only has to be done once. For fine alignment, we run a VI that automatically scans thetrapping beam across the detection beam; first in a line along x and then along y. As the trapping
beam moves, it takes the trapped bead with it, and the bead scatters light from the detection beam
(Figure 2.11A). Thus, at every scan point we have (1) the real-space distance (in nm) of thetrapping beam from its start point, and (2) the QPD x or y voltage signal of the detection beam.
Due to the shape of the detection beam and bead, this signal takes the form of a derivative of a
Gaussian, an antisymmetric function (Figure 2.11B). The center point of this function occurs
when the bead is in the center of the detection beam (for a given scan axis). We therefore fit the58

Figure 2.11: Procedures for positioning and calibrating a bead.
A) The beam centering procedure using a trapped bead. The trapping laser (orange) is moved
through the detection laser (red) with bead in tow, via a PZT mirror. B) Example trace produced
from the procedure. The x QPD signal (black) is fit (green) to find the position difference
between the signal's center point and the scan's center point. The off-axis signal (y; red) is also
shown. There should be little crosstalk. C) Setting the bead height above the surface. Using thestage, the surface hits the trapped bead, displacing it, then is brought back down. D) Example
height curve. The point of contact between surface and bead is when the signal changes fromoscillating to sharply rising. E) A calibration curve for turning QPD voltages into position
measurements. The data (black) is fit (green) by a 7th-order polynomial (equation inset).

59
data to find the position offset (in nm) of that center point, then instruct the trapping beam tomove to that location. In this way, we center the trapping beam over the detection beam.
With the bead’s (and trap’s) xy position set, we next set its z position (i.e., the height ofthe bead above the surface). This z position will determine the amount of viscous drag on thebead as it diffuses within the optical trap, due to the frictional interaction between the fixed glass
surface and the water. The closer the bead is to the surface, the larger the viscosity of thesurrounding liquid. Happily, this relationship is quantitatively given in Faxen’s Law (37), so as
long as we can accurately set the z position of the bead, we will know the surrounding viscosity.
For tethered beads, the z position also determines the relative contributions of the lateral andaxial trap forces to the total trap force (sections 2.4.3 and 4.1.2).
We set the height by moving the sample chamber surface in z relative to the optical trap
center. However, this motion is not completely independent. Recall that the lasers are focused byan objective lens directly before they enter the sample chamber (through immersion oil).
Changing the position of the sample chamber affects how close or far it is from the objective
lens, and therefore how focused the laser beams are when they enter the chamber. Thus, as thesurface moves relative to the objective lens, the absolute position of the beams’ foci in spacechanges (Figure 2.11C). Luckily, the relationship between these motions has been previously
worked out (103). The trap center follows the sample chamber’s position in a simple 0.2x
proportional relation; if the sample chamber rises in z by 10 nm, then the trap will rise in z by 2
nm. Thus, when we want to change the distance between surface and trap center, we simply tell
the stage to move 1.2x the desired distance.
To accurately set the height, we scan the PZT stage in z upward a set amount: enough forthe surface to contact the bead, begin pushing it out of the trap, and then move back down to a

60
noncontact position. From that, we get an 810 nm QPD z voltage signal (Figure 2.11D). The
signal point where the overall behavior changes (from weak oscillations to smoothly increasing)is where the surface first comes into contact with the bead. Once we have this absolute stage
position (in nm), we can easily set the height. We decide what distance we want between thesurface and the bead (typically 200-400 nm), and tell the stage to move down 1.2x that distancefrom the contact point.
With a bead fully positioned in space, our last task is to calibrate the 810 nm QPD
voltage signals so that we know the real-space position they represent. This calibration procedure
is very similar to the beam centering procedure. We use the 1064 nm PZT mirror to scan thetrapped bead in x and y, and record the voltage response from the 810 nm. However, instead ofjust fitting the response with a derivative of a Gaussian to find an offset, we fit it with a 7th-orderpolynomial and extract eight calibration coefficients (Figure 2.11E). Now, whenever we collect a
QPD voltage signal Vx, we simply multiply powers of it by the eight coefficients to get the realspace position x. With this, the bead is positioned and calibrated.

2.4.2 Trap calibration and linearity
Now that we can measure bead position, we next want to measure the force on the bead. Within
the linear region of the optical trap, this force is simply 𝐹 = 𝑘T 𝑥, where 𝑘T is the trap stiffness
and 𝑥 is the bead position. Therefore, we need to determine two quantities: the trap stiffness in
the linear region, and the extent of the linear region. We calibrate the stiffness semi-frequently
because it depends on the intensity and focus of the trapping beam, which can change over time
due to laser and optics drift. We check the linearity rarely because, once the linear region isdetermined, we typically perform experiments far away from the edges of the region so that61
linearity is assured. Specifically, the linear region is almost always larger than ±60 nm from trap
center along any axis, and we try to keep the bead within ±40 nm (the trap stiffness calibration
methods let the bead passively diffuse in the trap, reaching ±5 nm at most).
We calibrate trap stiffness using the equipartition method and the power spectrum
method. Since the data collection process for both methods is the same (they differ only in theiranalysis), we collect a single data set, perform both analyses, and compare the results to ensure
consistency. The first step in a 𝑘T calibration is to trap, position, and calibrate a bead. Once thatis done, the 1064 nm AOM servo reference voltage VAOM is set to a low value (0.25 V), whichcorrespondingly lowers the trap intensity. The diffusion of the bead inside the trap is recorded fora time (typically for ~1.1 s at 120 kHz; Figure 2.12A), then VAOM is increased by some linear
increment. This process is repeated eight times, so that there are nine bead diffusion data sets
associated with nine VAOM values. The idea in this calibration is to derive a trap stiffness value
from each diffusion data set and match it to its corresponding VAOM value. A line is then fit to the
𝑘T vs. VAOM values, and its coefficients extracted (Figure 2.12B). With these coefficients, we cantransform any desired 𝑘T value into a VAOM value and send it to the AOM servo board. Since thetrap stiffness can differ in the x and y directions, we calibrate both directions and pick a single
one to use in any given experiment.
To calculate 𝑘T using the equipartition method, we use the fact that a system in thermal
equilibrium with some bulk reservoir has an average thermal energy in each degree of freedom1equal to 2 𝑘B 𝑇, where 𝑘B is Boltzmann’s constant and 𝑇 is the temperature. The potential energy
imparted to the bead by the optical trap counts as one of these degrees of freedom. As mentioned
1

in the introduction, the optical trap forms a harmonic trapping potential, expressed as 2 𝑘T 𝑥 2 ,where 𝑥 is the distance between the bead and trap centers along some axis, and 𝑘T is the trap

62

Figure 2.12: Trap stiffness calibration and linearity.
A) An example trace of bead diffusion while trapped in the optical trap. The data was takenusing a 750 nm diameter bead, until 131072 = 217 points were collected (120 kHz for ~1.1 s).
Data sets in powers of 2 aid the FFT part of the power spectrum analysis. B) The trap stiffness
vs AOM set voltage calibration curves for the optical trap's x-axis, obtained via the equipartition
method (cyan) and power spectrum method (magenta). As expected, trap stiffness increases
linearly with beam intensity (proportional to set voltage), and both methods agree well. C) The
power spectrum of the bead diffusion data in (A), across the analyzed bandwidth. D) A dragmeasurement of a 750 nm diameter bead along the y axis of the trap. Trap stiffness was set to
0.2 pN/nm, and the drag coefficient was 9.67e-6 pN(s/nm). Trap linearity can be clearly seen,
and begins to start falling off near ± 60 nm. Acronyms: AOM (acousto-optic modulator), FFT(fast Fourier transform).

63
1

stiffness along that axis. The average potential energy is therefore 2 𝑘T 〈𝑥 2 〉, where 〈𝑥 2 〉 is thebead’s position variance. This quantity can be easily calculated from the bead diffusion data, and𝑘B and 𝑇 are known. Therefore, we can calculate the trap stiffness 𝑘T .
The power spectrum method calculates 𝑘T differently. We first take the power spectrumof the diffusion data using a Fast Fourier Transform (FFT). In rough terms, taking the power
spectrum of a time signal (i.e., the diffusion data) decomposes the signal into its sinusoidal
components (its Fourier series), extracts the coefficient of each component (their relative
strength), and then plots those coefficients as a function of the Fourier component frequencies;
an example for a trapped bead can be seen in Figure 2.12C. Thus the power spectrum, true to its
name, shows the relative “power” of each component in the frequency spectrum of a time signal.
The power spectrum of a trapped bead has a nearly Lorentzian shape, and can be analytically fit
to extract Lorentzian parameters (104). The parameter of interest here is the power spectrum’s
corner frequency 𝑓0 , the frequency at which the power is reduced to 1/2 the thermal baseline. The
trap stiffness can be calculated from this frequency as 𝑘T = 2𝜋𝛽𝑓0 , where 𝛽 = 6𝜋𝜂𝑓 𝑟 is the dragcoefficient for the bead (𝜂𝑓 is the dynamic viscosity of the solution corrected by Faxen’s law(105, 106) and 𝑟 is the bead radius). Since both 𝜂𝑓 and 𝑟 are known, the viscous drag on the beadcan be calculated, and so the trap stiffness can be derived.
Note that the trap stiffness partially depends on the size of the bead (section 1.1). As there
is some size variation in our bead stock (usually ~3% in diameter), there will always be someerror in using any one stiffness calibration for a number of different beads (46). This is typicallynot a problem, as absolute force measurements are not a primary concern in my work. However,for experiments where extreme force accuracy is desired, we typically calibrate the trap stiffnessfor each bead used (via equipartition, since power spectrum requires an explicit radius input).

64
Note also that these methods cannot be used to derive the trap stiffness along the z axis, since wecannot calibrate the QPD z voltage into real-space position (without having to manually scan thetrapping beam focus through the detection beam). However, this is not a problem, as we only𝑦need to know the axial-to-lateral stiffness ratio (𝑘T𝑧 /𝑘T𝑥 or 𝑘T𝑧 /𝑘T ) (section 2.4.3). This ratio canbe calculated by performing a power spectrum analysis on uncalibrated bead diffusion data in z
and x (or y), and taking the ratio of the Lorentzian fit corner frequencies multiplied by the ratioof the axially corrected drag coefficients (106). When using r = 375 nm beads, this ratio is
typically 0.5; when using r = 250 nm beads, it is typically 0.33.
To measure the extent of the linear region of the trap, we perform a viscous drag assay. In
this assay, we trap, position, and calibrate a bead, then move the sample chamber at some fixedvelocity along the x or y axis using the PZT stage. As the bulk liquid in the chamber moves, adrag force tries to drag the trapped bead along with the liquid; however, this force is counteredby the trapping force. An equilibrium is reached at some position away from trap center, suchthat the trapping force 𝐹 = 𝑘T 𝑥 balances the drag force 𝐹 = 𝛽𝑣, where 𝛽 is the drag coefficientof the bead and 𝑣 is the velocity of the chamber relative to the stationary trap. The position 𝑥 is
measured and recorded, along with the corresponding velocity 𝑣. This is repeated at manydifferent velocities, both positive and negative, as the magnitude of 𝑣 is gradually increased.
When completed, we plot equilibrium position vs. sample velocity, as seen in Figure 2.12D. In
the linear region of the trap, 𝑘T is constant and so there is a linear relation between position andvelocity. At the end of the linear region, that relation starts to break down as 𝑘T starts to changewith position (seen at ±60 nm in Figure 2.12D). Thus we determine the linear region.

65
2.4.3 DNA tether centering and calibration
Now that bead force and position measurements can be made, only one more quantity needs tobe found before we can study DNA tethers in the trap: the location of the DNA molecule’s otherend, the surface-DNA attachment point. Figure 2.13A shows the basic geometry of a tethered
bead in a surface-coupled optical trap; when an experiment begins, the surface-tether attachment
point is located an unknown distance 𝑥stage away from the trap center. Once that attachment
point is known, geometric calculations can be performed (Figure 2.13B) to convert it, as well as
the bead position and force, into the biophysical quantities we care about: the force and extensionof the DNA molecule (5). That knowledge can then be used to quantitatively stretch and relax
the DNA tether by controllably moving the attachment point relative to the trap using the stage.
We find the attachment point by centering it directly under the trap using a stretchingprocedure. This completes two tasks at once, since finding the attachment point in this manneralso fully relaxes the DNA molecule (minimize tension) and readies the system for further
experiments. The DNA centering procedure is identical to the beam centering one in concept: wescan something along the x and y axes, collect antisymmetric signals, and fit them to find theirposition offset values. Here, we use the PZT stage to move the unknown attachment point alongeach axis. The process is as follows: we assign the initial PZT stage position an 𝑥stage value of 0
and calculate its resulting DNA force and extension (F-x) values. Then we move the stage acertain amount, update the 𝑥stage value, and calculate the F-x values again. This is repeated until
10 pN is measured for the DNA force, then the process is repeated in the opposite direction until–10 pN is measured (Figure 2.13C). The position of zero force between the positive and negative
force portions when the DNA-surface attachment point is directly underneath the trap. We findthis position by fitting the signal with a two-sided worm-like chain (WLC) function which has

66

Figure 2.13: Surface-coupled geometry calculations and DNA centering.
A) Cartoon of the surface-coupled geometry used in this optical trap. The bead is pulled out of
the trap center by the surface-bound DNA tether. B) Equations for various parameters ofinterest, given the geometry of (A). Note that these relations only hold true within the linear
region of the trap. C) A two-sided, force-extension curve used for centering the DNA under the
trap. Data (black) is fit (green) to a two-sided WLC curve. The fit reports the persistence length,
extension offset, and contour length.

67
the zero-force offset value as a parameter. This process is performed on both axes iteratively tocenter the DNA tether under the trap (often two times is enough to center to ~1 nm accuracy).
The WLC fit also allows us to simultaneously perform a rough calibration of the DNA tether to
see whether it has the expected WLC DNA parameters. Specifically, we fit for the DNA tether
contour length 𝐿c and persistence length 𝐿p . If these values are roughly correct, we know thebead is connected to the surface by a single, undamaged DNA molecule. If either value issignificantly off, the bead is connected by multiple DNA molecules and/or damaged molecules.
In that case, we stop the procedure and move on to a new tethered bead.
Note that the diagram in Figure 2.12A is roughly to scale, and demonstrates why we use
DNA lengths of 3000–6000 bp (~1–2 𝜇m at contour length 0.338 nm/bp). Since the axial-tolateral trap stiffness ratio is roughly 1/2, the trap is twice as strong in the lateral direction as theaxial one. To keep the bead firmly in the linear regime of the trap, we want the majority of thetrapping force to come from this lateral force, which means we want to stretch the DNA tether
nearly horizontally (small angle between surface and tether).

2.4.4 Overstretching assay
With the tethered bead system fully positioned and calibrated, biophysical experiments can now
be performed. One of the most common and useful experiments I run is the overstretching assay.
This assay dynamically stretches and relaxes a DNA tether, often through its overstretching (OS)transition. The assay is dynamic in that F-x data is collected while stretching takes place (i.e.,
while the PZT stage is moving), not before or after. In contrast, the DNA centering procedure
only collects data after the PZT stage finishes moving. This dynamic nature introduces sometiming challenges into the LabVIEW code, as discussed below.

68
At the start of the assay, we increase the trap stiffness from 0.2 pN/nm to 0.8 – 1 pN/nm,
since we want to stretch the tether to a high force but still remain within the linear region of the
trap. Unfortunately, increasing the trap stiffness (laser intensity) by so much so suddenly causes
the objective lens to thermally expand. This thermal expansion moves the focus of the laser
beams, so that the trap center (and thus bead) increase in z position relative to the sample surface.
This increase in height, if not accounted for, can make tethers appear shorter than they actuallyare. This problem can be post-processed out, or mitigated by pre-warming the objective lensbefore the start of the assay. The trap stiffness should not be set higher during the centering andcalibration procedures, since the DNA tether can be damaged. Specifically, 1064 nm light caninteract with the polystyrene bead to produce reactive oxygen species, which can damage DNA(107). The production rate increases with laser intensity, so we try to only use intense beams
when the bead is not directly above the whole biomolecule; this way reactive species are more
likely to diffuse away than interact with the DNA molecule.
With the trap stiffness set, a long, concatenated GPIB command string is created and sent
to the PZT stage controller. This string contains the full stage movement plan for the entire
assay: move a set distance at a set speed and either (1) end, or (2) wait and pull back the same
distance at the same speed. After this string is sent, the VI starts recording bead position andforce data, as well as calculating where the stage should be at each time step. Note that we do notmeasure stage position during this dynamic assay; we infer it from the planned motion as acontrol parameter. The only time when this becomes a problem is during the wait between theforward and backward stage motion. Since this change in motion involves a lot of acceleration,
extra time needs to be added to the GPIB command string to account for the delay imposed by
the mechanical limitations of the stage’s acceleration. Once the stage finishes its command,

69
calculated stage data and measured bead data are transformed into DNA F-x data. The raw andprocess data signals are then saved, along with all assay parameters.

2.4.5 Active force-clamp assay
A force-clamp assay continuously measures the extension of a DNA molecule while
keeping the force on the DNA fixed to some value. Keeping the force fixed in this manner
ensures that DNA extension changes are solely due to biological processes, and are notconvolved with mechanical stretching from force changes. Both passive and active force-clamp
procedures exist (26, 108). We typically use active force clamps in this lab, and have two
methods: either the force is clamped by modulating the trap stiffness or stage position. Active
force clamps work much like the laser and stage servo loops: a reference force is set, DNA
tension is continuously measured, and if the current force differs from the reference force, thetrap stiffness or stage position is changed to bring the current force back toward the reference.
The benefits of using trap stiffness as the clamp variable are its speed and ease of use dueto the 1064 nm AOM servo loop, resulting in a high loop bandwidth (~1-10 kHz). It can also beeasily paired with active stage stabilization to further improve detection accuracy. The costs arethat the clamp is limited to the trap stiffness’s small available range; prolonged force changes
eventually cannot be stopped if they require the trap stiffness to exceed this range. Thermal noise
also gets added to the system due to the constantly varying trap stiffness, which makes the clamp
work even harder. Finally, the trap stiffness in this clamp obviously cannot be set to a stable,high value, which can be very useful for briefly reducing noise (see section 4.1.2). The benefitsof using stage position as the clamp variable are its large available range of motion and lack ofadditional noise. For all systems under study here, stage-based force clamps cannot run out of

70
range. The trap stiffness is also a free parameter in this setup, allowing for brief noise reduction.
The costs are its slower overall bandwidth (~100 Hz), due to the fact that changing the stage’s
position takes much more time than changing the trap stiffness.
To begin a force-clamp assay, the PZT stage is moved in rapid steps until the DNA force
reaches the reference force (usually 8 pN). If active stage stabilization is used, the 850 nm beamis then centered over a nearby fiducial mark and calibrated so that its QPD voltage outputs canbe read as positions. The force-clamp loop then begins to keep the DNA force within ±0.05 pNof the reference force. During each iteration of the clamp loop, force and extension data arecalculated from raw data and both (raw and processed data) are saved to disk. This process is
continued until the loop is manually stopped by the user. If a stage-based clamp is running, thetrap stiffness can be manually changed at will, which can be useful during high-resolution
experiments as a way to increase signal-to-noise (see section 4.1.2).

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Chapter 3
Force-activated DNA substrates

In this chapter, I detail the work performed on force-activated DNA substrates. First, I introduce
the rationale for creating such substrates, including how they can improve certain SMFS
experiments (section 3.1). Second, I describe two force-activated substrates that we designed,
constructed, and validated: a construct that yields 1002 nt of 50% GC ssDNA when activated,and one that yields a 20 bp DNA hairpin and 33 nt of ssDNA when activated (section 3.2). Third,
I go through the biophysical experiment we used to demonstrate the efficacy of these constructs:
a single-molecule measurement of the on-rate of the E. coil RecQ helicase using DNA hairpin
substrates (section 3.3). The work described in these three sections can also be found in Ref.
(109). Lastly, I detail the more advanced substrates we have made, including 120 bp DNA
hairpins with distinct runs of high and low GC content meant to improve %GC-dependent
unwinding velocity measurements (section 3.4).

3.1

Introduction to force-activation

SMFS experiments that study protein–nucleic-acid complexes typically rely upon dsDNA as a
substrate, and are often initiated by adding a critical component (e.g. nucleotide triphosphates)(110). However, a wide array of proteins, including motor proteins such as translocases and72helicases, may not bind directly to dsDNA, but instead bind to ssDNA and/or to ssDNA/dsDNA
junctions. Unfortunately, preparing substrates containing sections of ssDNA within longer
dsDNA molecules can be an inefficient process, which hampers their widespread application,
particularly for optical-trapping-based assays that require micrometer-scale dsDNA.
An important innovation in high-resolution SMFS helicase studies was the developmentof the hairpin-unwinding assay (33). In this assay, a long dsDNA molecule containing a short
hairpin and a small single-stranded nucleic-acid region is stretched (Figure 3.1A). The singlestranded region is typically slightly larger than the helicase’s footprint, so that the helicase binds
within a few nucleotides (nt) of the hairpin. Unwinding activity is detected via length changes inthe taut DNA substrate. For every 1 base pair (bp) of the hairpin unwound, 2 nt of ssDNA are put
under tension. Compared to directly measuring helicase motion along dsDNA (1 bp = 0.34 nm),the hairpin-unwinding assay provides a 3-fold mechanical amplification since the contour lengthof the taut DNA increases by 1.12 nm (2 × 0.56 nm/nt) (3, 50). Although initially applied to an
RNA helicase (33) and the ribosome (34), such assays have been extended to studies of DNA
helicases using both optical traps (111, 112) and magnetic tweezers (101, 113). Unfortunately,
the standard protocol for making these substrates involves a three-way hybridization (Figure
3.1B, left) (33), typically a laborious and inefficient process in our hands and others (114, 115).
Such assays are additionally complicated by the relatively low throughput of high-resolution
optical traps (42) and by fluid flows necessary to initiate unwinding (e.g. introducing ATP).
An alternative way to generate ssDNA is to mechanically denature dsDNA using force.
In the first experiment to demonstrate purely force-induced dissociation (i.e. no added chemical
denaturants), an atomic force microscope (AFM) was used to rapidly stretch the DNA (v = 0.15–
3 μm/s) (53). The resulting dissociation of a full ssDNA strand occurred at high force (120–250

73

Figure 3.1: Introduction to force-activated DNA substrates.
A) A cartoon of a single-molecule hairpin-unwinding assay using a surface-coupled optical trap.
Inset: a DNA hairpin is unwound by a helicase that initially binds to the small segment of
ssDNA adjacent to the hairpin. B) Left: the standard method to create ss-dsDNA substrates forsuch assays is via a three-way hybridization. The small arcs represent ligase molecules. Right: aforce-activated substrate contains a pair of enzymatically induced, site-specific nicks located on
the same strand. Small triangles represent nicking enzymes. Stretching this dsDNA molecule
partway into the overstretching transition induces dissociation. C) A force-extension curve for a2-um-long dsDNA molecule shows the overstretching transition at 65 pN as a function of thenormalized extension (nm/bp). Reproduced from Okoniewski et al., 2017 with permission from
Oxford University Press.

74
pN, depending on v) relative to DNA’s overstretching transition (65 pN) (3, 4). Recall that DNA
overstretching is characterized by a 70% increase in dsDNA extension at 65 pN (Figure 3.1C)
and often occurs by force-induced ssDNA dissociation starting from free ends or nicks,
commonly referred to as ‘peeling’ (58, 60, 61). However, previous experiments that pulled
dsDNA into and even just past the overstretching transition did not lead to its dissociation into
two complementary strands (3, 4, 50, 58-61, 116) (except when the DNA was ≤ 30 bp long(117)). Strand stabilization during and after overstretching is speculated to arise from islands ofrepeated GC base pairs (54) that suppress local DNA melting and peeling (58, 60, 61). Thisconjecture explains the much higher forces required to induce strand dissociation in the rapid
pulling assay (53, 54) and more recent results where sustained force above the overstretchingtransition was needed for dissociation (75 pN for ≥ 5 s) (116). This latter protocol, we note, didgenerate the controlled release of large segments of ssDNA defined by a single site-specific nick.
Despite this success (116), several important issues remain in such substrates: (i) high sustained
forces can lead to tether failure, (ii) force-activation efficiency decreases under ionic conditions
that stabilize base-pair formation (e.g. 150 mM NaCl), and (iii) more sophisticated DNA
structures, such as those needed for hairpin-unwinding assays, have not been realized.
To overcome these limitations, we developed two force-activated DNA substrates. In thefirst substrate (Figure 3.1B, right), a pair of enzymatically introduced nicks were separated by 1
kbp. The 50%-GC sequence between these nicks had no adjacent GC base pairs and, as a result,dissociated when we stretched the construct partway into the overstretching transition.
Importantly, the construct contained no ssDNA until activated, providing user-controlled
initiation of protein–ssDNA interaction at the single-molecule level. We designed a second DNA
substrate to leverage this capability so that we could study helicases using the hairpin-unwinding

75
assay. Specifically, we sequentially activated individual molecules of this hairpin-unwinding
substrate to measure the on-rate (kon) of the E. coli RecQ DNA helicase binding to ssDNA,
traditionally a difficult parameter to measure in a SMFS assay (101).

3.2

Validation of two force-activated DNA substrates

In the next two subsections, I detail the design, construction, and validation of two force
activated DNA substrates: one which activates to reveal 1002 nt of exactly 50% GC ssDNA(section 3.2.1), and one which reveals a 20 bp tetraloop DNA hairpin and 33 nt ssDNA bindingsite for hairpin-unwinding assays (section 3.2.2). The construction of these substrates follows thegeneral scheme laid out in section 2.3.1, with important differences pointed out as needed. The
substrates were validated using the overstretching assay described in section 2.4.4. Detailed
protocols on substrate construction can be found in the supplementary material of Ref. (109).
We analyzed the resulting force-extension curves by fitting the total extension as a sumof dsDNA and ssDNA extensions, using the Odijk eWLC for dsDNA (5, 118) and the extensible
freely-jointed-chain (eFJC) for ssDNA (3). Our goal was to determine the amount of ssDNA
generated after force activation. To constrain the fits, we fixed the dsDNA component of thetotal extension (using 0.338 nm/bp and the expected number of base pairs in the construct). This
was necessary for accurate fits, since the dsDNA component was so much larger and would
dominate the fits if left variable. We then fit for the ssDNA extension in both the initial and postactivation curves, which yielded ssDNA contour length values. The difference between the fitted
ssDNA contour lengths yields the number of ssDNA nt created, while also removing any
absolute offsets due to variability in bead size and linker length. In this process, we fixed thepersistence length (Lp) and stretch modulus (S) for ss and dsDNA (ssDNA: Lp = 0.75 nm and S =76
800 pN; dsDNA: Lp = 45 nm and S = 1200 pN). For the hairpin construct, we also took into
account the hairpin’s finite width of 2 nm. The reported uncertainties are the standard deviationsin the fitted parameters as calculated by the fitting routine (WaveMetrics, Igor Pro 6).

3.2.1 1002 nt 50% GC substrate
We hypothesized that efficient strand dissociation at 65 pN could be promoted by engineeringout GC islands. To do so, we developed a 1002 bp target sequence that had 50% GC content but
no adjacent GC base pairs (i.e. strictly alternates between GC and AT base pairs). The exact
sequence was designed using custom software that produced one hundred ten-bp segments byrandomly selecting bases to insert into each AT or GC spot. The full sequence was then run
through an autocorrelation routine to ensure no large sequence complements existed that could
yield stable secondary structure formation. For this proof-of-concept demonstration, we chose a
1000-bp segment so that we balanced the cost of synthesis with investigating a sufficiently long
DNA segment that would exhibit expected overstretching characteristics. Dissociation of this
1000-bp target sequence was facilitated by a pair of enzymatically induced, site-specific nicks(via restriction enzymes Nt.BspQI and Nb.BsmI), as shown in Figure 3.2A. The recognition site
for Nb.BsmI added 2 nt to the 50%-GC sequence. This full sequence was ordered from Genscript(Piscataway, NJ, USA). To facilitate optical trapping, GC-clamp sequences and traditional
endonuclease recognition sites (PflmI and BbsI) were added so that this designed sequence could
be ligated into a larger construct.
This larger construct, consisting of a 1667 bp ‘left handle’ and a 3670 bp ‘right handle,’
was assembled from sections of the M13mp18 plasmid. Briefly, we individually polymerase
chain reaction (PCR) amplified and purified the left handle, right handle and 50%-GC segments.

77

Figure 3.2: Forced dissociation of a 1002-nt DNA segment at ~65 pN.
A) Substrate schematic showing the 1002-bp 50% GC segment, nick sites, GC clamps, and 5'end labels. Nt.BspQI and Nb.BsmI are enzymes that introduce site-specific nicks. B) A set ofthree sequential force-extension curves at 2 um/s pulling velocity shows hysteresis in thestretching cycle. However, no permanent ssDNA strand dissociation is observed whenstretching the DNA molecule only 30% of the way through the overstretching transition (black).
The stage motion was reversed immediately to relax the DNA to zero force (cyan), and thenstretched again (magenta). C) A similar set of three force-extension curves, but with successfulforce-activation (identified by the permanent contour length changes observed in the cyan andmagenta traces). ssDNA dissociates when the construct is pulled 50% of the way through theoverstretching transition. The thick gray curve is a representative full overstretching curve.
Reproduced from Okoniewski et al., 2017 with permission from Oxford University Press.

78
The PCR products of the left handle and the 50%-GC sequence were then cut at their PflmI
endonuclease sites, combined together at an equimolar ratio and ligated. The purified ligatedproduct and the right handle were then cut with BbsI and ligated together. We then cloned thisfully ligated construct into a bacterial plasmid. The plasmid was transformed into competent E.
coli, grown up, and isolated. The isolated plasmids were verified by sequencing. Importantly, theabove process only had to be performed once. Once the full construct was in a plasmid, it was
efficiently produced for single-molecule application via a single PCR followed by site-specificnicking, without the need for any ligation. Specifically, for our optical-trapping application, weused primers that were 5’-labeled with biotin and DBCO, similar to earlier work (99). The
resulting purified and 5’-labeled PCR product was then nicked sequentially with the restriction
endonucleases Nt.BspQI and Nb.BsmI.
To demonstrate force-activation, we used a pull-relax-pull overstretching assay (section
2.4.4) where the stage was moved at v = 2 μm/s. By varying the predetermined stretching
distance, we controlled how far into the overstretching transition the DNA was extended (Figure
3.2B,C). As expected, the initial force-extension curve (black) for F < 40 pN was well fit by aneWLC model for dsDNA with contour length equal to that of the full construct (Figure 3.3). Thisagreement shows that the nicked DNA was fully double-stranded prior to force activation.
By pulling our engineered DNA construct halfway through the overstretching transition,
we achieved essentially instantaneous dissociation of a 1002 nt ssDNA segment. Notably, rapid
force-activation of this substrate required pulling the construct a minimum distance into theoverstretching transition. For small (≤ 30%) excursions into the overstretching transition, we
observed the canonical hysteresis between the stretching and relaxation force-extension curves
for such a dynamic assay (3). This hysteresis was caused by peeling of ssDNA from free ends or79

Figure 3.3: Quantifying force activation in the 50% GC construct.
Force-extension curves before (black) and after (magenta) force activation. The initial curve wasanalyzed using an eWLC model using a fixed dsDNA contour length (2179.8 nm = 0.338 nm/bp
× 6,449 bp). The fit (green) to the fully dsDNA construct returned a “ssDNA contour length”
value of 32.0 ± 0.3 nm, which accounted for variation in bead size variation and linker lengths(biotin-streptavidin and silane-PEG-azide-DBCO). The force-extension curve of the activatedsubstrate was fit the same way using a fixed dsDNA contour length of 1841.1 nm (0.338 nm/bp
× 5447 bp). This second fit (cyan) returned a ssDNA contour length of 589.6 ± 5.1 nm. The
difference in contour length was 557.6 ± 5.1 nm. Using a ssDNA contour length of 0.56 nm/nt,
this yields 996 ± 9 nt of ssDNA exposed during activation. This finding is in excellent
agreement with the expected value of 1002 nt. For consistency, both fits were performed from15 to 35 pN to avoid the low-force secondary structure unfolding seen in the activated substrate.
Model eWLC curves were extended to 0 pN for presentation using the determined parameters.
Uncertainties represent best fit ± s.d. Reproduced from Okoniewski et al., 2017 with permission
from Oxford University Press.

80
nicks, but such peeling intermediates quickly reformed fully duplex DNA once the force wassufficiently reduced (Figure 3.2B, cyan) (58, 59). On the other hand, when the construct was
stretched 50% into the overstretching transition (Figure 3.2C), a permanent increase in contour
length was observed between the initial and subsequent force-extension curves, a clear signal for
activation. To determine the amount of ssDNA generated, we compared WLC fits for the initialand second stretching curves (black and magenta, respectively). This analysis yielded 996 ± 9 nt
of ssDNA (best fit ± SD), in quantitative agreement with the 1002 nt ssDNA expected from ourconstruct (Figure 3.3). Importantly, every DNA construct that completed the more extensive
stretching cycle was activated (N = 22).
The observed 30% maximum distance into the overstretching transition before permanent
strand displacement suggests that peeling from free ends was concurrent with the force-induced
dissociation of the target strand at v = 2 μm/s. This interpretation is based upon previous work,
which showed peeling from free ends or nicks is required for hysteresis while hysteresis is notobserved in a topologically closed but rotationally free substrates (59). To be quantitative, this
30% excursion is 2-fold larger than that needed to overstretch just the 1002-bp nicked region.
Hence, we expect that there is peeling from the free ends of our construct as well as the sitespecific nicks. Such peeling had no adverse effects in this construct, but extra GC clamps could
be added to suppress it, if desired.
Beyond achieving efficient generation of ssDNA, these substrates also prove theconjecture that GC-islands lead to strand stabilization during and after overstretching (54).
Unlike extensive previous studies that used natural sequences containing GC islands, oursubstrate had exactly a 50% GC content with no adjacent GC base pairs. Upon stretching, thisengineered substrate still exhibited the standard overstretching transition at 65 pN, but we now

81
observed dissociation of a large strand (1 kbp) halfway through the overstretching transition.
Overall, this result, in conjunction with aforementioned 20 years of previous studies on natural
DNA sequences, shows that GC islands are critical to strand stabilization. Moreover, we expectthat strand dissociation within the 65 pN plateau generalizes to arbitrary lengths of 50% GC
segments above a minimum threshold of ∼20–30 bp (117), though maximum distance into theoverstretching transition before permanent stand displacement should vary.

3.2.2 Hairpin-unwinding substrate
Having established our ability to rapidly dissociate large segments of ssDNA, we next focusedon forming the more complicated structure needed for hairpin-unwinding assays. To do so, we
designed a substrate for a 3’-to-5’ DNA helicase that consisted of a 33 nt binding site adjacent to
a 20 bp hairpin stem capped by a 4 nt loop. We based the hairpin and binding site sequences on apreviously published hairpin (20R50–4T) (20) and previously published oligo (called oligo #3 inRef. (74)), respectively. Encoding the hairpin into the substrate was a technical challenge, since a
20-bp hairpin within a PCR primer is undesirable (Figure 3.4A). We solved this problem byencoding each half of the hairpin stem, as well as the loop region, into separate PCR primers.
Each primer also had one nick site for the endonuclease Nb.BbvCI, and one primer contained thesequence for the helicase landing site. Critically, the restriction site for the endonuclease BstXI
was encoded within the loop region of each primer (Figure 3.4B). Each primer was then used tocreate a separate PCR product, which we referred to as the ‘left’ and ‘right’ halves. The BstXI
sites on both molecules were then cut, yielding complementary, non-palindromic 4-nt ssDNA
overhangs. The left and right halves were then annealed and ligated (Figure 3.4C).

82

Figure 3.4: Challenges and solution to the 20-bp hairpin construct.
A) Cartoon depicting the challenge of using a PCR primer that fully encodes a hairpin.
Specifically, undesirable intra- and inter-strand hybridization will form. Such behavior will
negatively impact the subsequent PCR assays. B) To solve this, the hairpin is divided in halfusing the endonuclease BstXI. Its non-palindromic recognition sequence contains an internal 6bp that accommodates any sequence, and leaves a 4-nt overhang after cleavage. By using a nonpalindromic sequence (TTTT) in this region, self-annealing is avoided. C) Sketch of the steps
and sequences used to yield a DNA construct containing an internal hairpin. The two DNA
molecules encoding the two halves of the hairpin are nicked and then ligated together. Note, thishairpin sequence contains three instances of two adjacent GC base pairs, which did not interferewith force activation. Reproduced from Okoniewski et al., 2017 with permission from Oxford
University Press.

83
The resulting final construct was inserted into a plasmid for subsequent use as a PCR
template. We expect that the formation of a stable intrastrand hairpin within the template strand
during PCR extension (~70 °C) will limit the length of the hairpin stem that can be produced bythis strategy without additional post-PCR enzymatic steps. To improve the strength of thecoupling between the DNA and the optically trapped, streptavidin-coated beads, our forward
primer contained 4 biotins spaced every 9–10 bp. The reverse primer contained a 5’-DBCO. The
20-bp hairpin had 50%-GC content and the 33-nt section had 58%-GC content. We engineered
GC clamps into the shorter dsDNA handle to suppress unwanted fraying. The final construct had
a total size of 5529 bp (~1.8 μm) (Figure 3.5A).
Since the nicked strand contained only 83 nt, we first wanted to verify that the OS
transition was still required for activation. We repeated the rapid stretch-relax-stretch protocol
with a peak force of 55 pN, a high force just below the overstretching transition (Figure 3.5B).
No hysteresis was observed. Hence, no peeling occurred. Next, we repeated the stretch-relaxstretch protocol but stretched the construct just into the OS transition (7%). In this case,hysteresis and a contour length increase in the force-extension curves was observed (Figure
3.5C). Importantly, folding and unfolding transitions consistent with hairpin formation andrupture were now observed (Figure 3.5C, inset). As expected at v = 2 μm/s, there was now
hysteresis in the force-extension curves at low forces due to the non-equilibrium folding andunfolding of the DNA hairpin, similar to pioneering studies of RNA hairpins (17). We note that
substrate activation was successful in 100% of attempts (N > 300) at two relatively high ionicconcentrations that stabilize base pair formation (150 mM NaCl or 50 mM NaCl + 5 mM
MgCl2). To verify that the proper structure was formed, we analyzed segments of the second
stretching curve prior to and after hairpin unzipping (Figure 3.6). This analysis yielded 45 ± 1 nt

84

Figure 3.5: Force activation of a DNA hairpin substrate.
A) Schematic of the substrate with a more detailed view of the region around the strand to be
displaced. Enzymatically induced nick sites (Nb.BbvCl), GC clamps, and 5'-end labels shown.
B) A set of three force-extension curves at v = 2 um/s from a stretch-relax-stretch cycle shows
no hysteresis when the maximum force was kept just below the overstretching transition (black,cyan and magenta, respectively). C) A similar set of three force-extension curves but showing
force-activation when the construct was pulled just 7% of the way into the overstretchingtransition. Inset: a magnified view of the force-extension curves shows hairpin folding (cyan)
and unfolding (magenta) after activation. As expected, these curves span the initial forceextension curve of the fully dsDNA construct (black). Reproduced from Okoniewski et al., 2017with permission from Oxford University Press.

85

Figure 3.6: Determining the size of the activated hairpin.
A force-extension curve of the activated construct (red) shows a rupture of the hairpin at ~15
pN. The lower (2-15 pN) and upper (17-35 pN) sections were both fit using the same method
described for Figure 3.3. Both fits (green) used a fixed dsDNA contour length of 1840.75 nm =0.338 nm/bp × 5446 bp. In addition, the first fit includes a 2-nm offset to account for the
diameter of the DNA hairpin. These fits returned ssDNA contour lengths of 26.2 ± 0.4 nm and51.6 ± 0.2 nm before and after hairpin unfolding, respectively. The contour length difference
was 25.4 ± 0.5 nm, which corresponds to 45 ± 1 nt using a 0.56 nm/nt ssDNA contour length.
This result quantitatively agrees with the expected value of 44 nt. Uncertainties represent best fit± s.d. Reproduced from Okoniewski et al., 2017 with permission from Oxford University Press.

86(best fit ± SD), in agreement with the expected 44 nt introduced by hairpin rupture. Hence, we
generated a valid substrate for the hairpin-unwinding assay via force activation.

3.3

Single-molecule measurement of E. coli RecQ on-rate

To demonstrate how force-activated substrates can enhance helicases studies, we measured therate at which a protein binds to its target, the on-rate (kon), for the E. coli DNA helicase RecQ
binding to ssDNA. Determining on-rates is traditionally difficult in single-molecule assays (101),in part because the time associated with buffer exchange and flow-induced mechanicalperturbations degrades high-resolution assays. Our force-activated substrate assay removed theseissues, since we could start with a pure dsDNA construct––our inactive hairpin substrate––
stretched to 8 pN and premixed with helicases and saturating ATP [i.e. 1 mM ATP versus KM =16 μM (101)]. By rapidly activating the substrate (v = 5 μm/s), we can cleanly measure the time
t1 between activation and the start of unwinding (Figure 3.7A) using a 8 pN force-clamp assay(section 2.4.5). As expected, for all measurements unwinding was only observed after activation.
The time t1 is the sum of the time it takes the helicase to bind to the newly created 33 nt ssDNA(ton) and the time it takes the helicase to translocate along ssDNA to the dsDNA hairpin andbegin unwinding (ttrans). By measuring at saturating ATP but low enzyme concentration, we
made ttrans small relative to ton (~0.3 s vs. ~10 s, as the maximum expected ttrans ≈ 19 nt / 70 nt/s)(72, 95). Hence, our measurement of t1 yields ton, which in turn yields kon = 1/(τon[E]), where [E]is the concentration of RecQ and τon is the mean of the ton distribution).
We used E. coli RecQ DNA helicase in this study since RecQ is well characterized andcan bind to an internal segment of ssDNA prior to unwinding dsDNA (65). Backgroundinformation on RecQ helicases can be found in section 1.3. To focus on RecQ’s helicase activity,

87
we used a truncated version of RecQ for this assay, RecQΔH, which lacks its final domain, thehelicase-and-RNaseD–C terminal (HRDC) domain (77). In comparison to wild type RecQ,
RecQΔH shows no change in processivity nor mechanochemical coupling (93), but removal of the
HRDC domain increases ATPase activity by a factor of 2, and weakens binding to ssDNA by afactor of 2–5 (i.e., increased KD) (72). The buffer conditions used [30 mM Tris-HCl (pH 7.5), 50
mM NaCl, 5 mM MgCl2, 0.4% Tween-20, 1 mM DTT, and 1 mM ATP] are common for RecQ
family helicases, and can be traced back to an early study of human BLM (119).
To determine kon, we measured t1 at two enzyme concentrations that were much lower
than RecQΔH’s KD for ssDNA (90 nM) (72). In this limit, t1 (and thus ton) should scale inverselywith enzyme concentration. Specifically, we measured 70 individual molecules at 200 pM and
67 at 100 pM, all in a single day. Histograms of t1 were well fit by a single exponential andyielded average values of τ1 = 13.7 ± 0.7 s and 26.3 ± 1.8 s (best fit ± SD) for 200 and 100 pM,
respectively (Figure 3.7B). Importantly, our results agreed with the expectation that τ1 at 100 pM
should be twice τ1 at 200 pM. The dependence of t1 values on RecQ concentration coupled withthe exponential distribution of t1 at both concentrations shows that the dsDNA hairpin was
unwound by a RecQΔH monomer rather than a dimer, an ongoing question for RecQ (87, 90) andother helicases (120). Further analysis––assuming t1 ≈ ton––yielded kon = 3.7 ± 0.6 × 108 M−1s−1(best fit ± SD), where the reported uncertainty includes both the uncertainty in the fit as well as
enzyme concentration. This value for kon is near the ‘diffusion limit,’ the maximum rate at which
a molecule undergoing a random walk in 3D can collide with its target (121).
Our single-molecule determination of kon for RecQΔH is in close agreement with the one
prior measurement from an ensemble stopped-flow assay (95). In that study, 54-nt segments of
ssDNA were used as targets, in contrast to the 33-nt segment in the present work. Our value for

88

Figure 3.7: Force-activation enables an efficient single-molecule on-rate assay.
A) An extension vs. time trace prior to and immediately after force activation of the hairpinunwinding substrate in the presence of RecQΔH and saturating ATP. After a time t1, unwindingof the newly created hairpin led to an increase in extension when measured under constant load
(F = 8 pN). Note, the decrease in extension after activation matched the expected value (10 nm).
Activation occurred in ~0.5 s (orange bar). B) The normalized histograms of t1 at two different
enzyme concentrations well below the KD of RecQΔH for ssDNA. An exponential fit to thehistograms yielded the average time, denoted τ1. All the data were acquired in a single day [N =70 at 200 pM and N = 67 at 100 pM]. Reproduced from Okoniewski et al., 2017 withpermission from Oxford University Press.

89
kon is ∼1.5-fold larger than the previous determination (see supplemental material of Ref. (109)for in-depth calculation). Potential causes for this difference––albeit a relatively small one for a
kinetic parameter––include the taut conformation of our ssDNA and the fact that it is flanked bylong dsDNA handles, which could facilitate the diffusional search process (121).
One way our force-activated hairpin-unwinding assay can be improved is by decreasing
the activation time (currently 0.25 s). The fundamental limit for this time is set by the round trip
time of the stretch-relax cycle. This limit varies with stretching velocity and distance traveled by
the stage motion. For the present construct held at 8 pN, the round trip distance was 1000 nm.
Shorter constructs would decrease this time and are also advantageous since high-resolution
assays benefit from stiffer linkers. In addition, future work is needed to fully investigate thevelocity dependence of force activation, as we found that a small percentage (30%) of the 50%GC substrates failed to activate when pulled halfway into the overstretching transition at 20
μm/s, whereas all of the substrates activated at 2 μm/s.

3.4

Advanced substrates and %GC-dependent unwinding velocity assay

With the concept of force-activated substrates well-demonstrated, we decided to create moreadvanced versions of the hairpin substrate. The first improvement we focused on was increasing
the stem length: 20 bp is sufficient for many assays, but longer stems are often better. However,
we did not want to increase the length so much that the PCR reactions would start failing. We
decided that doubling the stem length to 40 bp would be a good first step. Following the processlaid out in section 3.2.2, we generated this new construct. Happily, increasing to 40 bp stems waseasy; no reduction in PCR efficiency or end product formation was observed, and all new
substrates force-activated successfully. So, in the same vein, we also generated a reverse version

90of this construct for 5’ to 3’ helicase unwinding. This simply involved switching the location of
the ssDNA landing site from downstream of the hairpin (for 3’ to 5’ motion) to upstream of it.
As expected, this construct worked just as well as its sister substrate, and both became our new
standard, general-use substrates for helicase studies. With the hairpin stems successfully
extended, we next pursued whether the GC content of the stem could be increased while still
maintaining successful activation.
To that end, we designed two 20 bp hairpins with increased GC content, one with 66%
GC and one with 75% GC. To our mild surprise, both could be made and successfully activated.
With these substrates, we intended to perform RecQ unwinding assays to measure %GCdependent unwinding velocities and compare our results to previously published values (122).
Unfortunately, this proved difficult, as RecQ often does strange things when within ~8 bp of thebase and top of hairpins (see sections 4.2 and 4.3). For a 20 bp hairpin, this leaves little room forclean unwinding. We thus decided to increase their stem lengths to 40 bp, as above. After testing,it was clear that the 40 bp 66% GC substrate activates successfully while the 40 bp 75% GC doesnot. The obvious explanation is that this length and %GC is too much for the overstretchingtransition to easily peel away, which is not entirely surprising, since the GC clamps we use to
explicitly prevent peeling in other parts of the substrate are around this length and GC content(~30 bp of 80% GC DNA, c.f. Figure 3.2A).
As development on the above constructs progressed, we designed a more advanced forceactivated hairpin for unwinding velocity measurements: a hairpin that has multiple, distinct
sections of GC content within a single stem. This substrate would improve unwinding velocity
measurements in several ways: (1) it would enable faster data collection since multiple %GC
regions are contained in a single substrate, and thus a single data trace; (2) the data from those91
different GC regions would be directly comparable in the clearest way possible, since they areplainly from the same molecule in the same buffer conditions; (3) and as before, it would enable
controlled data collection since the substrate must be activated. The second benefit is of special
note, since many enzymes possess static heterogeneity, meaning that any given batch of the
enzyme can have several sub-populations which behave differently from each other, such as
having different catalytic speeds or binding properties (this is likely due to the sub-populations
having slightly different folded structures) (123, 124). Using a substrate that can internallycompare the %GC-dependent unwinding of single members of each of these sub-populations
could be a great aid to data clarity.
Using our knowledge from previous constructs, we designed these substrates using 40 bp
building blocks. Specifically, the main construct has three 40 bp sections in a row: the initial andfinal 40 bp of DNA have 50% GC content, while the middle 40 bp has 0% GC (all AT basepairs). In addition, a control construct was designed with 50% GC all the way through (Figure
3.8A,B, top). Generating these 120 bp force-activated hairpin substrates was more complicatedthan prior constructs, since they could not be put into a single plasmid; the longer hairpin would
be nearly impossible to melt during a normal PCR reaction. To accommodate this, the order of
events was slightly changed (c.f. Figure 3.4C). Instead of ligating everything together andinserting it into a single plasmid, the two hairpin halves (and their associated DNA handles) were
each placed into plasmids and PCR amplified. The final hairpin ligation step at the loop was thenperformed after all PCRs were completed. While this process makes the construction slightly
harder and less efficient, it did successfully produce the substrates.
Once the 120 bp hairpin constructs were gel purified, we tried to activate them using thesame kind of pull-relax-pull overstretching assay as before. Noticeably, not all of the molecules

92
could be activated. Some tethers just did not activate, no matter how many times and how far
into the OS transition they were pulled. In clear contrast, the molecules that did activate usually
did so quickly and easily. While the cause of this disparity is not fully understood, the simplest
explanation is that the DNA nicking was not completely efficient, leaving some molecules witheither one or both of their restriction sites un-nicked (gel assays support this interpretation).
Roughly 25% of the molecules tested from both the 50-0-50% and 50-50-50% batches could not
be activated. While suboptimal, this only made the process of the data collection a little slower,
as non-activated molecules were quickly abandoned for new ones. For the tethers that did
activate, their F-x curves look as expected for both types of substrates (Figure 3.8A,B, lower).
For the 50-0-50% substrate, the 120 bp hairpin unzips in two distinct steps: the first 80 bp of thestem open at once, then the final 40 bp (confirmed by contour length fits). On the other hand, thefull 50% GC hairpin does not have multiple distinct pops, but instead smoothly unzips until
reaching a final barrier which is popped open.
With the substrates made, we next sought to demonstrate their previously listed benefits
in a helicase unwinding velocity assay. For this, we used the 50-0-50% substrates and the
RecQΔH helicases used in section 3.3. Prior measurements of how RecQΔH unwinding velocity
varies with GC content have been made (122), and we sought to reproduce these findings within
a single substrate. To do this, we used saturating ATP (1 mM) and recorded RecQΔH unwinding
the hairpin (Figure 3.8C). The full unwinding of 120 bp roughly equals a total extension changeof 90 nm in our 8 pN force clamp; 30 nm per 40 bp section. We chose to break up this total
extension change into 10 nm sections for analysis. As stated above, RecQ can do strange things
while near the base and top of a hairpin. Therefore, the first and last 10 nm sections of an
unwinding trace were excluded from analysis. For a fully unwound hairpin, this left four 10 nm

93Figure 3.8: Design and application of 120 bp force-activated hairpin substrates.
A) Top: Diagram of the 120 bp hairpin substrate containing 40 bp of 50% GC, 40 bp of 0% GC,
and 40 bp of 50% GC. Everything else was kept similar to the 20 bp hairpin design. Bottom:
Force vs. extension traces of the hairpin's successful activation. The hairpin was found to zip /unzip in two sharp jumps due to the 40 bp of 0% GC. B) The same information, but for the 120
bp hairpin containing only 50% GC. Here the hairpin was found to zip / unzip in one sharp jump
and one continous annealing / peeling event. C) Example trace of RecQ-dH unwinding the 50-050 %GC hairpin at saturating ATP. Each GC region corresponded to 30 nm of extension change
while held in an 8 pN force clamp. D) Histograms of average unwinding velocities along 10 nm
stretches of the 0% GC (red, N=470) and 50% GC (blue, N=823) regions. Both distributions
were fit with Gaussians and their mean velocities recorded. The ratio of the 50% GC mean
velocity to the 0% GC mean velocity is 0.83.

94

Figure 3.9: 66% GC unwinding velocities and validation.
A) Histogram of mean RecQ unwinding speeds on a 40 bp 66% GC hairpin. Data was analyzed
exactly like the histograms in Figure 3.8D. B) Plot of RecQ unwinding velocity ratios vs. %GC
content from a prior assay given in black. The 50% / 0% and 66% / 0% ratios collected withforce-activated substrates are plotted as magenta stars. Figure reproduced from Manosas et al.,2010 with permission from Oxford University Press.

95
sections of 50% GC unwinding and three sections of 0% GC unwinding. From each of these
sections, a simple linear slope was calculated (total extension change / total time). The resulting
velocity values were then binned into histograms based on GC content (Figure 3.8D; 0% GC
histogram N = 470, 50% GC histogram N = 823). These histograms were fit with Gaussians to
extract their mean values: 54 ± 0.5 nm/s for 0% GC, 45 ± 0.5 nm/s for 50% GC (best fit ± fit
error). The ratio of these two values is 0.83 ± 0.01, which agrees extremely well with thepreviously found value of 0.82 (Figure 3.9B) (122). As a final coda for this investigation, we alsocollected a data set of RecQΔH unwinding velocities using the 40 bp 66% GC hairpins, so that thehigher than average GC content substrates could be demonstrated as well. This unwinding data
was analyzed from two 10 nm bins (5-15 and 15-25 nm), but otherwise was collected exactly as
above. The distribution, while containing fewer points than the 0% and 50% GC data sets (N =94), still turned out Gaussian, and its fit produced a mean value of 40 ± 1 nm/s (Figure 3.9A).
The ratio of this unwinding velocity with the 0% GC velocity is 0.74 ± 0.02, again in excellent
agreement with the previously found value of 0.76 (Figure 3.9B).
Such excellent agreement with prior work shows that these advanced force-activatedhairpin substrates do function as intended, and can be used to enhance DNA unwinding assays.
As before, data was straightforward to collect, and most of the data sets were collected in single
day sittings. It should be noted that, while the unwinding velocity ratios agreed very well withprior work, the absolute values of the velocities differed by around 20%. For example, ourmeasured velocity at 0% GC was 54 nm/s, while in the prior work it was ~67.5 nm/s. However,this difference is to be expected, as the assay conditions differ slightly between our work andtheirs. The most prominent difference is the temperature difference between the two assays: ourunwinding took place at 22 °C, while in the prior work the temperature was 29 °C. Another96
difference is the force across the DNA during the unwinding: we used 8 pN while the prior work
used 9 pN. Both of these differences would lead to faster unwinding taking place in the prior
work, relative to ours (as observed).

97

Chapter 4
RecQ helicase unwinding

In this chapter, I explain the work done to examine the detailed unwinding behavior of the E. coli
RecQ helicase, given the establishment of force-activated DNA substrates and monomeric RecQ
unwinding in the last chapter. First, I detail this chapter’s major drive: the quest to resolve thefundamental physical step size of RecQ (section 4.1). I next present some of the interesting
behaviors that were observed while studying the step size: the ATP-less unwinding or “popping”
RecQ can perform (section 4.2) and some of the more exotic binding events RecQ can initiate,including rapid strand switching (section 4.3).

4.1

The physical unwinding step size

In this section, I cover the investigation into RecQ’s fundamental physical step size. First, I
review the information known about RecQ’s stepping behavior (section 4.1.1). Then, I explain
the many optimizations that must be done to achieve the signal-to-noise (S/N) needed for a
successful stepping assay, both to the trapping instrument (section 4.1.2) and the biochemicalsolution conditions (section 4.1.3). Finally, I present the final stepping results achieved in this
instrument: my collected data support RecQ having a 1 bp forward and backward step size, anddo not support the more exotic stepping models proposed by other papers (section 4.1.4).

98

4.1.1 Introduction to RecQ stepping
As stated in section 1.3, the exact mechanics of how RecQ performs its fundamental function,
dsDNA unwinding, are still unclear. Given that RecQ is an SF2 helicase with two RecA-likedomains, it likely has a physical translocation step size of 1 nt when moving along ssDNA,
analogous to other SF1 and SF2 helicases (68). However, this 1 nt physical translocation may notdirectly translate into 1 bp unwinding. As seen in Figure 1.4C, RecQ most likely keeps the ssdsDNA junction under tension, and pulls the connected ssDNA such that the duplex slides out of
the binding site and splits apart. The question is whether, for a given 1 nt translocation step, theduplex always unwinds 1 bp.
Bulk biochemical studies of RecQ have repeatedly found a chemical unwinding step sizeof 1 bp (71, 72, 87). In other words, they have found that, on average, 1 bp is unwound for every
1 ATP hydrolyzed by the enzyme. Note that this is only a bulk average; it does not say whether
or not 1 bp is unwound for each ATP hydrolyzed. Some translocation steps could result in no
unwinding, while others could result in multiple base pairs unwinding once the tension acrossthem is too great. Such a mechanism could explain the multiple RecQ unwinding step sizes
observed in a recent FRET study (92), although the paper itself offers a different interpretation:
they posit RecQ can occlude nucleotides as they are unwound, releasing them later. Supporting
this hypothesis, our collaborators at the NIH believe they have observed a 5 bp kinetic step sizefor RecQ due to occlusion effects (91).
Thus, the mechanics of RecQ’s unwinding remain unclear, and a high-resolution opticaltrapping assay could help illuminate them. Furthermore, given that RecQ is the representative
member of the RecQ helicase family, one of the major families within helicase superfamily 2,

99
uncovering the mechanics of how this helicase performs its basic function is of fundamental
interest. We therefore decided to investigate RecQ unwinding in high resolution, focusing on thetruncated version RecQΔH for simplicity, since its helicase properties are the same as the fulllength, wild-type enzyme, while its binding properties are much simpler (see section 1.3).

4.1.2 Optimizing optical trapping conditions
Obtaining 1-bp resolution in any stepping assay is a very difficult task. Almost every aspect ofthe measurement has to be optimized, since the expected signal from the enzyme step is so low
compared to the overall noise of the system (largely from thermal noise). In this section, I
explain the parameters that determine both the signal and noise in a surface-coupled opticaltrapping stepping assay, and what must be done to increase the signal and decrease the noise.
For a single DNA held under tension, the signal obtained from helicase unwinding will
be some change in both DNA force and extension. For reasons detailed later, extension is by far
the better signal to use. Since helicases unwind dsDNA into ssDNA, we should be able toquantify unwinding changes using the eWLC and eFJC, the models which predict the F-x
relationship of dsDNA and ssDNA, respectively (Figure 4.1A). Exactly how we implement thesemodels depends on the geometry of the DNA substrate used. For example, say we measured
RecQ unwinding with a gapped DNA substrate (such as the 1000 nt force-activated construct insection 3.2.1), where every 1 bp of dsDNA unwound leaves behind 1 nt of ssDNA under tension;
let’s set the tension to 8 pN. The expected signal for this process is then 0.376 nm – 0.322 nm =0.054 nm (adding 1 nt and subtracting 1 bp): in other words, half an Angstrom. That signal isvery small, so unsurprisingly gapped DNA substrates are not used in stepping assays. The bestgeometry currently available is the hairpin assay: unwinding 1 bp of the hairpin stem releases

100
two nt of ssDNA into the larger tether, while keeping the hairpin stem width fixed. At 8 pN, thisgives an extension signal of 0.75 nm, a 15x amplification over the gapped DNA geometry. Thisis the DNA substrate type used here, and the first optimization for increased stepping signal.
Substrate tension is the only other parameter that directly impacts the signal. As seen in
the F-x models, the higher the force across the substrate, the larger the extension values for both
dsDNA and ssDNA. Since larger ssDNA extension produces larger unwinding signals, highsubstrate tension is clearly desirable. However, there is an upper limit to force in a hairpin assay:
the force at which the hairpin unzips. If the tension across the hairpin reaches or even nears thatforce, the hairpin can immediately unzip and all further unwinding signal is lost. For the 20–40
bp stem hairpins detailed in Chapter 3, this opening force is ~15 pN. For safety, the highest force
across these substrates should be ~12 pN. Yet we cannot immediately pick that value as theunwinding force, because force also affects assay noise, not just signal. This choice is dealt withat the end of the section, after the other noise sources have been described.
That said, we can note that without a force clamp the tension in any unwinding assay will
constantly lower as new ssDNA is added into the tether, since adding more length produces moreslack. Moreover, the amount the force lowers by depends on the full tether length, and thuschanges after every unwinding step. These length-dependent, non-constant force changes are
why force is not used as the signal in unwinding assays. Furthermore, even minor reductions intether force can reduce the extension unwinding signal dramatically. When a 1 bp step releases 2
nt of ssDNA into the larger tether, it instantaneously reduces the tension across the rest of thedsDNA tether (~3000 bp), attenuating the overall extension increase. To get a quantitative
understanding of how, I created a simulation of a DNA hairpin substrate held under tension in asurface-coupled optical trap.

101

Figure 4.1: DNA mechanistic properties and step simulations.
A) The dsDNA eWLC model and the ssDNA eFJC model, both as functions of extension permonomer unit (bp or nt). B) Nucleic acid polymer stiffness curves as a function of force. Madefrom the derivatives of the curves in (A). At a given force, the DNA k value must be divided by
the number of monomer units in the polymer to get the true stiffness. C) Simulations of RecQ 1bp stepping on my hairpin substrate held in an 8 pN force clamp with a proportional gain = 1.0.
Each step creates an instantaneous, small drop in force that attenuates the extension signal
greatly. It also takes the force clamp ~100 ms to recover from this drop (with no noise present in
the system). This clearly demonstrates the need for a fast force clamp for accurate detection. D)
Same as (C), but with 1.0 pN/nm trap stiffness. Higher system stiffness creates larger force
drops and attenuation, but faster recovery.

102
In these simulations, I set a 3000 bp tether with a 39 nt binding site in an 8 pN force
clamp with a 250 nm radius bead. The stage position was the control variable, and data was
generated by solving the geometric constraint equations (see Figure 2.12) and F-x models. Forease of use, the generated data was assigned a 1000 Hz bandwidth; this is low enough to justify
the fact that all thermal equilibration takes place during a single data point, while high enough tosensibly run a simulated active force clamp (at 200 Hz with a proportional gain of 1.0). I thenassigned 2 nt of ssDNA to be added to the tether every 200 ms (equivalent to a 1 bp hairpinstep). Running this simulation at 0.2 pN/nm and 1.0 pN/nm trap stiffness produces some very
interesting results (Figure 4.1C and 4.1D, respectively). In the 0.2 pN/nm case, we see that 1 bp
steps reduce the tension from 8.00 pN to 7.94 pN, a 0.75% change. However, that is enough toreduce the initial extension change from the ideal 0.75 nm to 0.40 nm, almost a 50% change!
Again, this is due to the ~3000 bp in the handles all shortening just a little, attenuating the overall
extension increase from the addition of 2 nt. Moreover, with a gain of 1.0 it takes the force clamp
over 100 ms to raise the force back to 8 pN and the signal to 0.75 nm. That is clearly too slow,
and would end up smearing together the extension stepping signal in a real assay. Thus theclamp’s gain value is clearly an important parameter, which will depend on the force and thenoise in the system. Raising the trap stiffness to 1.0 pN/nm keeps the same behavior, except theforce drop is higher due to the higher overall system stiffness. This further attenuates the initialsignal, but speeds up the force-clamp correction due to the higher ΔF values. Further down, I
will discuss the importance of high system stiffness in more detail.
To summarize, the extension signal depends both on the clamped force value and how
well the force clamp keeps that value fixed. Force-clamp performance is key, as fluctuations inforce distort the extension signal. Thus optimizing the force clamp is another critical task. While103
using a full PID (proportional-integral-differential) servo loop may sound tempting, I found that
including integral and differential terms just adds noise to the extension signal. The clamp
becomes too complex and starts tracking noise or over-compensating for sporadic unwinding.
Therefore I only use proportional gain, and have to empirically optimize it for a given assay
geometry and set force.
With this, we have exhausted signal optimizations; everything else involves mitigating
noise sources, both in extension and force. These noise sources come in two types: thermal beadnoise and instrument drift. Thermal bead noise is a fundamental part of trapping experiments, sowhile it can be mitigated, it can never be completely removed. Instrument drift, on the other
hand, is not fundamental and can ideally be eliminated (although real instruments are non-ideal).
Instrument drift comes in two main flavors: beam drift (discussed in section 2.1.4) and surface
drift (discussed in section 2.1.5). I will first examine thermal bead noise, as its optimization is theunderlying reason for multiple design decisions.
The power spectral density of a trapped bead’s thermal motion in solution can bemathematically described by a Lorenzian distribution (a type of Cauchy distribution) (37):𝜎𝑥2 (Δ𝑓) =𝑘B 𝑇 Δ𝑓 𝑑𝑓∫,
𝜋 2 𝛽 0 𝑓 2 + 𝑓02(4.1)

where 𝜎𝑥2 (Δ𝑓) is the position variance of the bead as a function of measurement bandwidth Δ𝑓,𝑘B 𝑇 is the thermal energy of solution, 𝛽 is the bead’s drag coefficient, and 𝑓0 is the corner (orroll-off) frequency of the optical trap. Importantly, integrating from 0 to infinity produces theequipartition relationship 𝜎𝑥2 = 𝑘B 𝑇⁄𝑘trap , where 𝑘trap = 2𝜋𝛽𝑓0 is the trap stiffness (see section
2.4.2; the log-log plot of the integrand in Eq. (4.1) can be seen in Figure 2.12C). For a trapped
bead tethered by a DNA molecule, the expression gets slightly altered: the stiffness of the DNA
is added to the trap stiffness (as they are springs in parallel). DNA stiffness can be computed by

104
taking the derivative of the F-x curves in Figure 4.1A. The resulting functions (Figure 4.1B) canbe used to compute tether stiffness by taking the stiffness at a specific force and dividing it by
the number of bp or nt in the substrate. In other words, longer nucleic acid polymers are more
flexible, while shorter ones are stiffer.
With the DNA stiffness accounted for, we can compute the noise value we care about: thestandard error in the mean bead position. This quantity is the square root of 𝜎𝑥2 divided by the
number of independent data points averaged together from the bead position data. Roughly, theroll-off frequency of the system (trap + DNA) specifies how quickly these independent data
points are collected. The expression for this can be written as (125):

𝜎𝑥SEM

𝑘B 𝑇 ∆𝑓
𝑘B 𝑇 12𝜋 2 𝜂 𝑟 Δ𝑓
√𝑟 Δ𝑓
√
√
√
=
=
=
= 𝜋√12𝑘B 𝑇𝜂
,
𝑘T 𝑓0
𝑘T
𝑘T
𝑘T
√𝑁
𝜎𝑥

(4.2)

where 𝜎𝑥SEM is the standard error, 𝑁 = 𝑓0 ⁄∆𝑓 is the number of independent data points averaged
together (∆𝑓 is the final bandwidth, which must be lower than 𝑓0 ), 𝜂 is the dynamic viscosity ofthe solution, 𝑟 is the bead radius, and 𝑘T = 𝑘trap + 𝑘DNA is the total stiffness of the system.
Note 𝛽 = 6𝜋𝜂𝑟 for a spherical object in liquid. Also note that 𝑘DNA is ~0.2 pN/nm for my tether
lengths and set force, while 𝑘trap gets set to 0.2 – 1.0 pN/nm. The last expression in Eq. (4.2) has
been split into two terms for clarity. The first has constants and temperature-dependent inputs
that are not very variable; obviously the constants are fixed, and 𝑇 and 𝜂 do not change muchover a few Kelvin, the temperature range of my trapping assay. Therefore, this term is roughly
constant, and noise reduction efforts must be directed towards the second term.
At first glance, the bead noise seems to: (1) increase with bead size by √𝑟, (2) increase
with the measurement bandwidth by √Δ𝑓, and (3) decrease proportional to 𝑘T . Yet that analysis
ignores how the parameters interrelate. Recall from section 1.1 that the trap stiffness has a 𝑟 3

105
dependence (for beads with 𝑟 much smaller than ~400 nm, the beam radius) (46). Thus, for agiven fixed 𝑘T , larger beads will have higher noise. But for a given fixed laser power larger
beads produce a much higher 𝑘T , and thus reduce the overall bead noise.
𝜎𝑥SEM ∝

√𝑟
√𝑟
√𝑟
=
∝ 3
.
𝑘T
𝑘trap + 𝑘DNA 𝑟 + 𝑘DNA

(4.3)

One way to understand this relationship is to note how bead size and trap stiffness affect thecorrelation time 𝜏 = 𝛽 ⁄𝑘T , the average time interval between independent (i.e., uncorrelated)data points. Increasing the bead’s size increases the drag on the bead, which increases the timeinterval during which its movements are correlated. Increasing the system stiffness decreases thattime interval, since a stiffer system corrects the bead’s thermal vibrations faster. The best size
choice will depend on the exact optical setup, since the 𝑟 3 scaling falls off as beads get as large
as the beam width. Increasing bead size further increases drag linearly while initially keeping
trap stiffness constant, and then eventually decreases trap stiffness, as the beads become toolarge for the trapping laser to properly surround (46).
Given this information, it is clear that beads can be too small or too large for stepping
assays. There are other complications: the first is that bead size also affects force noise. To get an
expression for that, we can multiply the bead’s position noise given in Eq. (4.2) by 𝑘T :𝜎𝐹SEM = 𝑘T 𝜎𝑥SEM = √2𝜋𝑘B 𝑇 𝛽 Δ𝑓 .

(4.4)

From this, it is clear that while the bead’s position noise can be reduced by increasing 𝑘T , theforce noise cannot. Thus increased bead size only increases the force noise, which makes theforce clamp harder to run, which degrades overall signal quality.
The second complication is that optimizing bead size and system stiffness takes place in a
2D geometry. From section 2.4.2, we know that trap stiffness has lateral and axial components,

106
and that the axial one is typically 2x to 3x weaker than the lateral one. To increase overall
stiffness, one must therefore pull the DNA tether at a very shallow angle, so that the contributionof the lateral trap stiffness is maximized (Figure 4.2A). However, this cannot be done with large
beads and short DNA tethers, as both will geometrically force the pulling angle to increase.
Trying to reduce the pulling angle by decreasing the height h between the bead and surface alsocomes with a cost: the viscous drag on the bead increases due to the proximity of a fixed surface.
This increased drag is quantified by Faxen’s law correction factors to the dynamic viscosity 𝜂 inboth directions (Figure 4.2B) (106). Increasing drag at fixed stiffness increases the correlation
time 𝜏 = 𝛽 ⁄𝑘T of the bead’s motion (i.e., lowers roll-off frequency via 𝑘trap = 2𝜋𝛽𝑓0 ), reducing
my ability to average down noise. Thus there is a careful balancing act between choosing theright bead size, height, and tether length to achieve the ideal assay geometry for low noise work.
After iterative testing of bead sizes, heights, and tether lengths, I settled on a bead radiusof r = 250 nm, a height between surface and bead edge of h = 200 nm, and a tether contour
length of 1000 nm (3000 bp of dsDNA). I found this combination, which produces an initialpulling angle of ~22°, to give the best mixture of noise characteristics and usability. In thistrapping instrument, beads with 250 nm radii can produce a trap stiffness of 1 pN/nm in thelateral direction when using slightly less than max laser power, sustainable for several seconds
before thermal drift starts to dominate. As seen in Figure 4.2B, a 250 nm radius bead at a heightof 200 nm above the surface induces a Faxen’s law drag correction of ~1.5x and 2.5x in thelateral and axial directions, respectively.
With tethering geometry optimized, we should further delve into the costs and benefits ofincreasing trap stiffness via the trapping laser power. This method of increasing 𝑘T is the mostcontrollable, since bead size and material are both fixed properties once the experiment begins;

107

Figure 4.2: Surface-coupled geometry optimization.
A) Cartoon depicting the geometry considerations that must go into designing a surface-coupledassay. The bead radius, height above the surface, and DNA length all affect the pulling angle θ
between the DNA and surface. The smaller the angle, the more the stronger lateral trap stiffness
contributes to the assay. B) Faxen’s law correction factors for the drag coefficient as a functionof height above the surface. The corrections factors for motion parallel to the surface (solidlines) are lower than those for motion perpendicular to the surface (dashed lines). In addition,
the larger the bead, the larger the correction factors at a fixed height.

108
laser power, on the other hand, can be rapidly changed thanks to the AOM servo loop (section
2.1.4). Increasing trap stiffness in this manner reduces the bead thermal noise dramatically, as
expected (Figure 4.3A), but comes with two costs: intense thermal drift in the instrument andpotential biological damage. Recall that the lasers pass through a large objective before entering
the sample plane (section 2.1.1). This objective, made up of multiple lenses glued into a single
metallic shell, is temperature sensitive. As laser light passes through it, the internal elements
absorb heat from the light and the entire piece expands. This thermal expansion distorts thebeams’ propagation, leading to laser drift in the sample plane. In tethered bead assays, thisproduces large drift effects in the extension trace (Figure 4.3B). Second, in addition to signal
degradation, more intense laser light can also harm biological molecules. 1064 nm light caninteract with polystyrene and molecular oxygen to produce reactive oxygen species, which
induce damage in both DNA and its linker chemistries (107). Oxygen scavenging systems canmitigate this damage (e.g., PCA-PCD, glucose oxidase, DTT), but higher laser light will
inevitably lead to higher rates of damage. Finally, higher laser light also heats the water near the
optical trap up a few Kelvin. However, as specified above, this temperature change is typicallynot enough to impact the S/N in a meaningful way.
By increasing the trap stiffness so dramatically, we soon run into a new noise floor. In
other words, we are able to reduce the bead thermal noise to such an extent that it is no longer
dominant. The subsequent dominant noise is, unsurprisingly, relative drift between the surface
and lasers. Even with active stabilization on both, small jerks around their mean positions canstill get through into the signal (Figure 4.3C). Given optimized servo loops, this noise cannot be
further reduced and sets the instrument’s noise floor. And unfortunately, this noise can be on thescale of the expected stepping signals. This means that not only is the time at high stiffness

109

Figure 4.3: Optimizing the trap stiffness.
A) The thermal noise in bead position can be significantly reduced by increasing the stiffness ofthe optical trap. Shown is 1200 Hz position data for a trapped r = 250 nm bead with either 0.2
pN/nm (grey) or 1.0 pN/nm (cyan) trap stiffness. The standard errors in the mean of these data
sets are 0.9 nm and 0.2 nm, respectively, giving an overall noise reduction of 4.5x, close to the
expected value of 5 for a 5-fold stiffness change (see Eq. 4.3). B) Going from low to high trap
stiffness by changing the laser power in the sample plane induces thermal drift. Shown is theextension data for an activated hairpin tether at 3 kHz (light) and 300 Hz (dark) at 8 pN. The
characteristic thermal drift for this process is a slight increase in extension, followed by a flat
period of little-to-no drift, followed by a rapid decrease in extension. This decrease willcontinue as long as the high trap stiffness is maintained, and will even continue briefly after thelaser power is decreased. These different types of drift are assumedly due to different parts ofthe microscope objective thermally expanding at different rates. Data is ideally collected in theflat region with no drift for maximum precision. C) Reducing the thermal noise via high trap
stiffness reveals the instrument noise floor. An activated hairpin tether held at 12 pN and 1.0
pN/nm will display instrument drift on the order of the desired stepping signals. This is the
major limitation of the assay (light: 3 kHz; dark: 300 Hz).

110
limited by thermal drift, we will also need multiple unwinding steps in a row in order to clearlydelineate between real steps and the noise floor. These demands on the experiment, combinedwith others detailed further below, will clearly reduce the throughput of useful data.
With the trap stiffness sorted, let us return to the clamped force value. As discussed
above, the benefits of high tension are an increased signal (to due higher ssDNA extensions) andslightly reduced noise (due to slightly stiffer dsDNA handles). Yet there is a cost to higher force:faster helicase unwinding speeds, since putting more tension across the dsDNA hairpin reduces
the energy needed to unwind it. Recall that resolving 1-bp steps through thermal noise involves
averaging down high bandwidth data to achieve better spatial precision at the cost of time
resolution. Ideally this averaging should only be done in between instances of helicase stepping.
If multiple steps get averaged into a single data point, obviously that stepping information is lostand all we get is a smear of increasing extension. Thus, the speed at which the helicase unwinds
sets a limit on the amount of data averaging, and thus the amount of noise reduction, actuallyachievable. That said, the effect force has on unwinding speed is enzyme-specific, and RecQ’s
speed has been shown to be nearly independent of force (making it a very “active” helicase)(122). This is at once a blessing and a curse: we can use high force to amplify the signal without
affecting the speed, but we cannot use low force to significantly slow down the helicase andthereby increase our noise averaging capacity. After many tests, I found that 8-12 pN worked
well, and that there is few observable difference between values in this range. Since earlier
magnetic tweezer assays on RecQ used 8 pN, that was my most commonly set force. However,
my later stepping work used 10 and 12 pN more frequently for slight increases to S/N.
As a final note, another way to potentially reduce RecQ unwinding speed is to lower thesolution temperature. Many enzymes have temperature-dependent activity, if thermal forces are

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critical for certain binding events, conformational changes, or oligomeric formations (recall theimportance of temperature in the oligomeric debate for RecQ in section 1.3). Unfortunately, thefew studies performed on the temperature dependence of RecQ’s unwinding activity have foundlittle change within my viable temperature range (85, 101). Given this, I did not pursue
temperature variation as a way to slow down RecQ.

4.1.3 Optimizing biochemical conditions
While we have exhausted optical trapping optimizations, there is still one major way to improve
the signal-to-noise ratio (S/N): slowing down unwinding speed. As stated above, the helicase
unwinding speed sets a upper bound on how much spatial averaging can be done before signals
get averaged together into a smear. Therefore, slow enzyme stepping is highly desirable, andsince RecQ requires ATP for its activity, the obvious way to reduce its speed is to reduce theconcentration of ATP in solution. By limiting [ATP], we make ATP binding rate-limiting in the
catalytic cycle, and then adjust that rate by adjusting [ATP]. This is a very common tactic in
stepping assays (26, 32), but in RecQ’s case there is a steep cost: RecQ’s off-rate from DNA, therate at which it unbinds and dissociates from the substrate, is also ATP-dependent (126). Higher[ATP] means RecQ is able to unwind more processively, staying on a single substrate for longer,
while lower [ATP] means it dissociates sooner, sometimes right after it binds. This introduces adifficult balancing act into signal resolution: on the one hand, we need the slow, well-resolved
steps that come from low [ATP]; on the other hand, we need enough consecutive steps toproperly active high trap stiffness for increased S/N and to discern the true step size of RecQ.
Since RecQ rapidly unwinds at saturating [ATP] (1 mM), finding an [ATP] value where
RecQ stepping can be resolved is a critical first step (Figure 4.4). As reported in section 3.4,

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Figure 4.4: RecQ processivity varies with [ATP].
Four traces of RecQ unwinding at 200 Hz bandwidth and 8 pN tension. The processivity of thehelicase lowers as [ATP] is lowered. The 1 μM ATP trace was collected at 1.0 pN/nm trap
stiffness for maximum precision, as opposed to the 0.2 pN/nm trap stiffness used in the otherthree traces.

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RecQ unwinds 50% GC DNA at an average speed of ~60 bp/s (45 nm/s) for 1 mM ATP and 8
pN of force. That gives a mean stepping frequency of one step per 16 ms, with most of the steps
occurring much faster (for an exponential distribution of stepping times). Since my active force
clamp runs once every 5-10 ms (depending on the exact assay), the clamp will not keep up andso will lose the steps in smear and noise. To slow down stepping, [ATP] must be reduced to nearor below its Michaelis-Menton constant for RecQ (the concentration at which RecQ’s ATPhydrolysis rate is half the saturating rate). Previous work found this value to be KM = 16 μM(101). Using 20 μM, 10 μM, and 1 μM [ATP] definitely slows mean RecQ unwinding, but also
shows the steep cost: processivity is lost and the steps are slowed down non-uniformly (Figure4.4). Even at low [ATP], there are often unwinding runs that have largely the same speed as
saturating ATP. Much of the slowdown arises from the decreased processivity, which introduces
not only the expected pausing but also apparent slippage into the traces. So while lowering[ATP] is helpful overall, the clear downsides provide motivation to try alternative methods for
slowing down RecQ.
Lower [ATP] slows RecQ unwinding because it extends the ATP binding step of thehydrolysis cycle (Figure 4.5A). However, that is not the only way to extend the binding step.
Another tactic for slowing down ATPases is to replace some portion of the ATP with nonhydrolysable analogs, which extend the ATP binding step by temporarily clogging the hydrolysis
pocket, forcing usable ATP molecules to wait until it becomes unclogged. This trick has thebenefit of keeping the ATPase in a conformation close to its ATP-bound state, extending thetime spent in that conformation instead of the ATP-unbound one. This is desirable for RecQ,
since it has higher affinity for DNA in its ATP-bound states (126). If I could slow down thehydrolysis cycle by using an analog while keeping the unwinding processivity high, I could

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better resolve RecQ’s overall stepping. Thus I spent time trying out different non-hydrolysable
ATP analogs to see if one of them could dramatically improve the assay.
However, adding in direct competitors to ATP is not automatically an improvement: if
certain criteria are not met, these analogs will just make the assay worse. For instance, to resolve
single steps, the analogs must on average interrupt the ATP hydrolysis cycle at least once per
cycle. It does no good if only one out of every dozen steps is delayed, as sequential 1 bp steps
will likely never been seen. The ideal analog should have a RecQ on-rate comparable to or
higher than ATP and a RecQ off-rate slower than the hydrolysis and product dissociation rate ofATP. This would enable the analog to out-compete ATP to RecQ and stall the enzyme, creating
the desired slowdown. Sadly, the binding affinity for any analog is likely to be lower than ATP’s
affinity (since ATP is the binding pocket’s sole target substrate), but increasing the relative
concentration of analog to ATP can compensate for this. The bigger sticking point is the off-rate:
we want an off-rate that is slow but not too slow. For example, if the off-rate is so slow than asingle step takes a minute, instrument drift can potentially obscure the steps, even with active
surface stabilization. Ideally, we would like an analog to slow the hydrolysis cycle down 2-foldto 10-fold. With these criteria in mind, I tested a variety of non-hydrolysable ATP analogs.
The analog I used first, and the most common non-hydrolysable ATP analog, is called
ATPγS. This molecule has the same structure as ATP except for one atom: the γ-phosphate’s
double-bounded oxygen is replaced with sulfur. This single swap renders it practically nonhydrolysable by many ATPases, including RecQ (74, 91). After running multiple unwinding
assays with [ATP] and [ATPγS] values ranging from 0 to 1 mM, I was able to confirm several
points about the interaction dynamics of RecQ, ATP, and ATPγS in solution. First, ATPγS has aslower on-rate and off-rate than ATP, as expected. For the on-rate, using an ATP:ATPγS ratio of

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Figure 4.5: RecQ interacts with non-hydrolysable ATP analogs.
A) A diagram for the typical kinetic pathway of RecQ's hydrolysis of ATP with approximate
rate constants. The rate constants are specified as either best-fits or limiting values (lower
bounds for k1, k-2, k3, and k4; upper bound for k-3. Associative rates given for pseudo-first-orderconditions). Figure reproduced from Sarlos et al., 2015 under the Creative Commons license. B)
RecQ unwinding trace with 50 μM ATP and 50 μM ATPγS. Data taken at 0.2 pN/nm trap
stiffness in 50 mM NaCl and 1 mM MgCl2. Light: 100 Hz, dark: 10 Hz. The pauses in this trace
were noticeably more uniformly distributed than other traces taken in these conditions. From
this, 1-bp forward and backward steps can be clearly seen, in addition to larger extension
increases. It is impossible to tell whether these larger increases are due to single multi-bp steps
or the smearing together of multiple 1-bp steps. C) Example traces of RecQ unwinding in thepresence of 0.5 mM ATP and 0.5 mM ApCpp (left), 0.5 mM AppCp (middle), or 0.5 mM
AppNHp (right). Very infrequent, very long pauses can be observed in all three traces.
Compared to ATPγS, these non-hydrolysable ATP analogs are therefore not as suited for aiding
1-bp stepping assays.

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1:1 does not provide enough reliable slowdown, while using a ratio of 1:5 completely stops all
unwinding activity. All available data indicate that ATPγS keeps RecQ bound to DNA while it isassociated with RecQ. For the off-rate, ATPγS seems to have a wide range of dissociation times
whose average is on the order of ~1 s. The best results from these mixed assays occur when ATP
is kept low, in the 10-50 μM range, to keep the overall RecQ speed low. I collected some bpscale stepping results with these conditions (Figure 4.5B), but overall the ATPγS-induced
slowdown still had many multi-bp steps and smeared unwinding runs. To see if I could obtainbetter-resolved unwinding, I tested several other non-hydrolysable ATP analogs to check
whether their kinetic rates were more in line with ATP.
The next analogs I used had additional atoms in their structures, in between the phosphategroups. If we denote ATP by Appp, where A is the nucleoside and the p’s are the phosphategroups, then these analogs were ApCpp, AppCp, and AppNHp, where the C’s are carbon
insertions and the NH is a nitrogen insertion. To test whether their kinetic rates were closer toATP’s than ATPγS, I ran RecQ unwinding assays with equal parts ATP and analog, using 0.5 M
for both to ensure that no pausing would arise from low [ATP]. The results of these tests showedthat all three analogs had much lower on- and off-rates compared to ATPγS (Figure 4.5C). Thus,
they were not viable replacements. At this point, it seemed that while doping the solution with
non-hydrolysable analogs was helpful, it would never provide the kind of uniform slowdown I
needed to ensure that all extension changes could be unambiguously designated as single signals.
With that determined, I then worked to see whether other steps of the hydrolysis cycle could be
extended, such as the actual hydrolysis step or product dissociation step.
To slow down the hydrolysis step, one would need to either mutate the RecQ or change acomponent in the hydrolysis reaction. These components include the triphosphate fuel source

117(usually ATP), the divalent cation that bridges the phosphate-protein interactions (usually Mg2+),or the salt and pH of the external environment (usually ~100 mM NaCl at pH 7.5). The overall
goal is to slightly distort the geometry of the RecQ hydrolysis reaction to make it less efficient,and thus slow it down. I initially tested environment changes, as they are easiest to implement;
they did nothing. Varying monovalent salt from 25 mM to 150 mM did not affect unwinding
speed in any measureable way. For the pH, prior bulk studies showed RecQ’s binding affinity for
ssDNA drops 2-fold at pH 5.5 and 8.5 (from a maximum at pH 6.0-7.5) (86). However, I havenever found a bulk measurement for how RecQ’s speed changes with pH. In my single-molecule
experiments, I did not observe any speed difference at pH 8.5, and was unable to measure RecQ
unwinding at pH 5.5 due to my beads adhering to the surface. From this, I determined thatvarying environmental conditions was not a fruitful path for slowing RecQ unwinding.
Another way to slow down the hydrolysis step is to replace ATP with a hydrolysable
analog. This allows the reaction to take place, but at a reduced rate since the analog does not fit
perfectly into the binding pocket. While investigating chemicals listed as non-hydrolysable ATPanalogs, I found that RecQ could actually hydrolyze one: ATPαS, the α-phosphate equivalent of
ATPγS. My initial experiments showed that RecQ unwinds ~2-fold slower with ATPαS vs. ATP(at 50 μM), but that most of the slowing is due to processivity loss and slippage, similar to the
speed decreases from lower [ATP]. ATPαS possesses two optical isomers, Sp-ATPαS and RpATPαS, and previous research found that multiple enzymes hydrolyze the Sp isomer while the Rp
isomer is rarely hydrolysable (this trend held for dATPαS versions, as well) (127-129). The firstbatch of ATPαS I tested was a 1:1 mixture of the two isomers, so it was unclear how RecQ
precisely interacted with each. To clear this up, we tested a pure stock of Rp-ATPαS, since priorwork implicated it as the less hydrolysable of the two (127-129).

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Running a RecQ assay with 20 μM ATP and either 0 or 100 μM Rp-ATPαS showed aclear processivity loss for all of the 100 μM traces; most could not even clear half of the hairpin
before dissociating (Figure 4.6A). This shows that the analog and helicase frequently interacted,
which is already better than some prior analogs (Figure 4.5C). Compared to the unwinding seen
with the 1:1 ATPαS mixture, this mixture has slower and less processive activity even with amuch higher overall concentration. Thus we can deduce that the Sp isomer in the 1:1 mixture is
the major component used in unwinding, and that RecQ interacts with Sp-ATPαS similar to ATP,akin to enzymes in past literature (127-129). We can therefore ignore the mixture and just use
ATP and Rp-ATPαS, or Rp-ATPαS alone (control experiments showed that RecQ can hydrolyze
Rp-ATPαS, although very poorly).
As a note, I found that environmental conditions affect ATPαS hydrolysis differently than
ATP hydrolysis. For ATP, RecQ unwinding was unchanged by monovalent salt variations; thiswas not true for the 1:1 mixture of ATPαS. Specifically, very little processive RecQ unwinding
was observed when using 150 mM NaCl (as opposed to the standard 50 mM), even with 1 mM
1:1 ATPαS. However, I found that the unwinding runs present at these conditions had the same
local speed as those at lower [NaCl], implying that the off-rate of the RecQ·ATPαS complex
from its DNA substrate is salt dependent, but that the binding and hydrolysis rates remain
unaffected. Sadly, that means this salt dependence cannot be used to slow RecQ.
For completeness, I also decided to check whether the ATP in RecQ assays could be
replaced by other NTPs, specifically GTP and UTP. One of the first papers on RecQ tested thehydrolysis activity of RecQ with GTP, and did not find any within the noise threshold (70).
However, because our lab had some GTP lying around, I double-checked it on the off-chance of
there being some very slow activity that could benefit my S/N. But as expected, there was no119
unwinding activity at 1 mM GTP. The UTP was much the same story, except the idea for it camefrom finding out that RecA can hydrolyze UTP poorly (130). Since RecQ’s hydrolysis pocket ismade up of two RecA-like domains, I chose to try it, as well. No activity was detected.
With the exception of mutation, there is only one more way to disrupt the hydrolysis
reaction: swap the Mg2+ that coordinate the ATP with different divalent cations. Investigating
how divalent-coordinated enzyme reactions change with divalent type is a standard procedure,
and was one of the first biochemical analyses performed on RecQ (70). In that work, the authors
found RecQ’s hydrolysis of ATP was most efficient with Mg2+, as expected. Swapping the Mg2+
for Mn2+, Ca2+, and Zn2+ reduced the hydrolysis efficiency to 96%, 89%, and 24% of the Mg2+
value, respectively. These findings tell us that in the RecQ binding pocket Mg2+ and Mn2+ act
interchangeably, with Ca2+ acting very similar. Zn2+ was the only ion exchange that significantly
reduced ATP hydrolysis efficiency, though the bulk assay could not say exactly how. Divalentswapping is especially popular for polymerase studies, since the kinetic changes induced by theswaps can be mapped to questions of direct biological interest, such as how metal contaminantscause mutations during replication (131). The cited review shows that these studies use a large
collection of divalent cations with multiple polymerases, and that they found some clear trends in
how the enzymes behaved with certain cations. I used this data and the work done previously on
RecQ to inform my choices for further divalent swapping.
In RecQ and the studied polymerases, swapping Mg2+ for Mn2+ did not change any
kinetic parameters, so I did not bother testing it. Since Ca2+ was in my lab already, I tried it as
my first swap to see what a 10% drop in bulk hydrolysis efficiency looks like in a singlemolecule assay. As expected, unwinding looked mostly the same, with the addition a few morepauses (Figure 4.6B). I next tried to repeat the same experiment with ZnCl2, but ran into the120

Figure 4.6: Slowing RecQ unwinding with ATPαS and divalent swaps.
A) RecQ unwinding traces with 20 μM ATP (black) and 20 μM ATP + 100 μM Rp-ATPαS(blue). 300 Hz traces taken at 0.2 pN/nm trap stiffness with 75 mM NaCl and 1 mM MgCl2. The
most striking effect of adding Rp-ATPαS is the processivity loss in all RecQ unwinding events.
The consistency of this loss implies that RecQ and Rp-ATPαS repeatedly interact during theunwinding traces. B) Comparison of RecQ unwinding at 1 mM ATP with either 1 mM Mg(black) or 1 mM Ca (orange) as the divalent cation. Other conditions similar to those in (A). As
expected from prior bulk ATP hydrolysis assays, RecQ can processively unwind DNA when
only Ca divalent cations are available. The only noticeable change in these single-molecule
traces was a slight increase in the number of pauses during unwinding. C) 1-bp steps resolved in
the presence of 1 mM Co divalent cations and 1 mM Rp-ATPαS. Light traces: 3000 Hz, dark
traces: 300 Hz. Data was taken at 1 pN/nm trap stiffness without DTT in solution, as sidereactions occur when it and Co are placed in Tris-HCl, NaCl solution. Left: At least 3 bp worthof unwinding occurs in a smear, followed by two well-resolved 1-bp steps forward and one 1-bp
step backwards. Right: Drift occurs that puts the unwinding trace slightly out of register withprior steps. However, four 1-bp steps forward can clearly be seen to occur after the backwards
step. Unfortunately, after this unwinding becomes a smear again.

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problem of zinc oxychlorides precipitating out of solution and causing my bead-DNA tether
complexes to adsorb to the surface (132). Replacing the zinc chloride with zinc acetate fixed thisproblem, only to show that while RecQ’s unwinding speed certainly drops when using Zn 2+, its
on-rate absolutely plummets. Acquiring just a few unwinding traces was a very time-consuming
task, and RecQ repeatedly slipped off of the DNA substrate, indicating that it was not wellbound. Since this may indicate Zn2+ is distorting the unwinding process too much (and thus notproviding biologically relevant information), I decided to try a few other divalents. First I tried
Sr2+, the next after Ca2+ in the alkaline earth metal column of the periodic table, but like in all
previous polymerase assays (131), it appears that RecQ too cannot use Sr2+ at all. This is due tosteric hindrance in some respect: either the ion cannot fit inside the binding pocket, or it issimply too large to correctly coordinate ATP for the hydrolysis reaction (133).
The next transition metal divalent I tried, Co2+, did work well. RecQ’s on-rate to DNA
with Co2+ was similar to its on-rate with Mg2+, and it processively unwound DNA while
simultaneously experiencing a ~2-fold unwinding slowdown. Because of its ease-of-use andeffective properties, Co2+ became my divalent of choice when not using Mg2+. As a note, I foundthat mixing DTT, CoCl2, and buffered solution with Tween-20 did not work well; a sickly brownsolution would form (DTT and CoCl2 mixed by themselves is fine). Since using a buffered
solution with Tween-20 is necessary when using polystyrene beads (or else they get stuck to theglass surface), I leave out DTT in all Co2+–based assays. To make sure this did not affect RecQ’s
activity in any significant way, I tested a side-by-side comparison of Mg2+-based RecQ
unwinding with and without DTT. There was no difference between the assays, meaning thatDTT acts solely as an oxygen scavenger in RecQ unwinding experiments (meaning its removal
will lead to more rapid DNA damage, but switching to new substrates can compensate for this).

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When combining Co2+ with ATP unwinding did not produce as much slowdown as I was
hoping, I decided to try it with 1 mM Rp-ATPαS, to see whether or not RecQ could hydrolyze
the molecule. Unwinding did occur, though with much lower processivity than with ATP (asexpected from the results of the prior ATP + Rp-ATPαS assay). With these conditions, I was able
to collect some bp-scale stepping runs, but again, with less processivity than I would like andwith most of the potential “steps” smeared together by fast unwinding (Figure 4.6C). With this, I
concluded the biochemical optimization process without a clear answer to a single, optimum
condition set. No conditions tested gave an unequivocally superior unwinding assay compared toall others; it was largely a give-and-take between speed, processivity, activity frequency, ease-ofuse, and binding strength. I therefore decided to summarize what RecQ stepping conclusions I
could obtain from these various condition sets.

4.1.4 Final RecQ stepping results
With all of this work done, what results did I obtain with my stepping assays? Unfortunately, I
never achieved sufficient resolution to unambiguously determine whether the few multi-bp steps
I observed were real or simply due to a few fast 1-bp steps being smeared together. This meansthat I cannot definitively comment on whether RecQ only takes 1 bp unwinding steps or whether
it can take multi-bp steps. With that said, the RecQ stepping data I did collect can be compared
to and contrasted with prior results, starting with the results of the 20 Hz FRET-under-tension
assay (92) that observed a wide distribution of RecQ step sizes.
In my clear stepping data, I primarily observed only 1-bp steps (Figure 4.7A). This data
mostly came from low [ATP], Mg2+-based traces, but also included a few traces from otherconditions (Co2+, Rp-ATPαS, ATP+ATPγS). I typically resolved runs lasting two to four steps;

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not enough for useful stepping statistics, but still plainly different from prior work (91, 92). On
that point, a number of 1-bp back steps were also resolvable (Figure 4.7B), which have neverbeen clearly shown for RecQ before and will be discussed further below. I was able to collectthese traces at forces from 8 – 12 pN and at [ATP] values from 1 μM to 1 mM, with clear steps
more common at lower [ATP] but still present at higher [ATP], especially when using RecQWT
instead of RecQΔH (the final HRDC domain is expected to alter stepping kinetics, but not thephysical step size). In fact, a large disparity in the speed of short stepping bursts exists (Figure
4.7C). To quantitatively analyze my stepping data, I took 50 clear stepping runs and used a stepfinding algorithm to determine step sizes. This algorithm was previously validated and publishedby my collaborators Keir Neuman and Yeonee Seol, and uses a t-test to determine significant
differences in extension data noise (101). My criteria for selecting “clear” stepping traces isbased on the visible presence of at least three extension dwells at regular intervals (for at least
two steps in a row), with no regard to the actual interval lengths. Thus no selection bias for a
specific step size was placed upon the data. With that, the step-finding algorithm found a total of
245 steps in the 50 traces. I then converted the calculated extension values into base pairs (basedon the applied force) and plotted the distribution (Figure 4.8A).
This distribution has a clear peak at 1 bp for forward steps, and a second peak at -1 bp forback steps. Other forward step sizes are present (notably at 0.5 bp), but the 1 bp size dominates
the distribution. This is in stark contrast to the previously published RecQ step size distribution
from the FRET-under-tension assay (Figure 4.8B). Here the authors, like me, only collected clear
traces with a few steps per trace, but instead of using a trap they used a FRET assay paired with a~6 pN intrinsically taut DNA substrate. The authors observed RecQ taking steps at 20 Hz withsizes ranging from 0.5 bp to 3.5 bp. Importantly, only ~15% of their recorded steps were 1 bp

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Figure 4.7: The E. coli RecQ helicase takes 1-bp physical unwinding steps.
All data collected using Mg2+, ATP, and 8 pN (for 0.75 nm Δx per 1 bp step). A) RecQ data wasanalyzed by eye for sequential, identifiable dwells (steps), with no criteria placed on spacing.
Almost all detected steps had 1-bp spacing. Most runs were limited to 2-3 steps due to noise orsmearing from rapid steps. B) 1-bp backsteps were regularly seen. Their speed was comparable
to forward steps in each assay, and occurred in RecQWT (left) and RecQΔH (middle & right). C)
Longer 1-bp runs at different time scales. Left: RecQWT sometimes produced good runs, but was
overall harder to work with. Middle: An exceptionally slow stepping trace. This plot and the one
above, taken on the same day, were the only time such slow steps could be collected. Right: A
fast, resolved stepping trace. The apparent 2-bp step is likely two very fast 1-bp steps.

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Figure 4.8: The high-resolution step size distribution strongly peaks at 1 bp.
A) A RecQ step size histogram collected using a t-test-based step-finding algorithm on my highresolution unwinding traces in my optical trap. A total of 245 steps were detected from 50
distinct unwinding runs. Data traces used in this distribution were collected at forces between 812 pN and [ATP] between 1 μM to 1 mM, then converted from extension into base pairs. The
underlying distributions for these conditions were all similar to the total distribution, showingthat the fundamental physical step size does not depend on force or [ATP]. B) The equivalent
RecQ stepping distribution from the FRET-under-tension assay. Data was collected on a ~6 pN
tensioned DNA substrate at 2 μM [ATP] and 20 Hz bandwidth. Figure reproduced from Ma etal., 2018 with permission from the American Chemical Society.

126(relative to the normalized counts of the rest of the distribution). In contrast, over 50% of theirsteps were either 1.5 bp or 2 bp (from a data set of 249 unwinding runs), with both individual
peaks being larger than the 1-bp peak. If this step size distribution was correct, I should haveseen it reproduced in my assay. But I did not. Instead, I found that ~80% of my forward steps
were clearly 1 bp, with the majority of the rest being either 1 bp steps broadened by noise orapparent combinations of 0.5 bp and 1.5 bp steps. This leaves two options: either (i) the multi-bp
step size distribution is wrong, and likely suffers from multiple fast steps blending together in the
20 Hz data, or (ii) my results are incorrect. Unsurprisingly, I think option (ii) is unlikely. Withmy high spatial and temporal precision, I was able to better resolve many fast unwinding runs as
being composed of successive 1-bp steps (c.f., Figure 4.7C right). At lower resolution, these runs
would have instead looked like larger, multi-bp steps. And if larger physical step sizes wereindeed more common than 1-bp steps, I should have seen them, as they would have been easierto resolve. But I did not. Therefore, I think it is likely that the multi-bp step distribution isincorrect: E. coli RecQ takes a majority of 1-bp unwinding steps, as expected from its known
translocation step size. That said, I cannot definitively state whether or not RecQ only takes 1-bp
steps; it is certainly possible that it rarely takes larger steps, and I certainly observed 0.5 bp steps
in my experiments, though my S/N is not high enough to further pursue that analysis.
Let us return to the subject of RecQ back steps during unwinding (Figure 4.7B). Theseback steps occurred both as single events and as multiple, distinct steps, typically once RecQ
made its way a few bp into the hairpin. Back steps were always soon followed by the resumptionof normal forward unwinding, evidence that the RecQ conformation during both step types is not
drastically different. In addition, the prevalence of back steps increased as [ATP] decreased.
Linking these observations with the hypothesized structure for RecQ binding to DNA (Figure

1271.4C) presents a possible mechanism: slippage of the RecA-like domains from their ssDNA
track. If the binding between the ssDNA nucleotides and the RecA-like domains briefly broke,
the RecQ zinc-binding and winged-helix domains that loosely associate to the dsDNA portion of
the junction could keep the protein loosely bound. In that instant, the ssDNA would be free fromtension and could start zipping itself. After one nucleotide zipped into the dsDNA, the remaining
ssDNA nucleotides would once again be in register with the RecA-like domains, allowing themto rebind. Given low [ATP] conditions, the process could repeat until the RecQ rebound to thesubstrate firmly. However, this mechanism does not adequately explain how multiple, relatively
slow 1-bp back steps can occur in succession. It is unlikely for slippage to repeat with such slow
regularity, often with a speed comparable to forward steps (c.f. Figure 4.7B).
Another possibility is that these back steps are really forward steps along the opposite
strand. In other words, RecQ briefly switches translocation strands and lets the unwound hairpin
close behind it as it moves away from the ss-dsDNA junction. It is clear from my data that RecQ
can switch strands in some capacity, as I often observe a single helicase unwind the same hairpin
multiple times. This behavior can be seen in prior figures (Figure 4.4, black and Figure 4.6A) as
well as in figures from upcoming sections (Figures 4.10A, 4.13A, and 4.14A). Such repeatedunwinding can only occur if, after unwinding the full hairpin stem, the helicase then continues
translocating down the complementary strand and eventually switches back to unwind the rezipped hairpin. If it is possible for RecQ to strand switch after unwinding the full hairpin, it may
be possible for it to strand switch during normal unwinding. This mechanism could explain boththe existence and dynamics of the observed back steps, but it is not without issues. The mostglaring issue is the large conformational change that needs to occur: can such a position switch

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happen quickly, without inducing large-scale hairpin zipping when the helicase lets go of thesubstrate (c.f. Figure 4.7B)?
To my knowledge, the E. coli RecQ has never been shown to take back steps before,
although strand-switch-based back steps have been observed in the A. thaliana RECQ2 homolog(134). It is not surprising that they went unobserved in previous E. coli RecQ studies, as bulk
assays could not detect them and single-molecule assays used either very high ATP (1 mM) orvery low ATP (~2-5 μM). Neither condition is ideal for producing back steps, since high [ATP]keeps RecQ driving forward with little chance of slippage or strand switching, while low [ATP]prevents RecQ from unwinding more than a couple steps. Without opened nucleotides to close,
and potentially enough room to strand switch, backsteps cannot occur or be detected.
In the course of this project, I have not seen any evidence of a 5-bp kinetic step, even in
longer stepping traces (Figure 4.7C). To review, my collaborators believe they see systematicpausing in RecQ unwinding once every ~5 bp, which they think arises from RecQ occluding andthen releasing unwound nucleotides (91). On the other hand, I have never observed this kind of
pausing, either in RecQΔH or RecQWT unwinding traces. To better understand this disagreement, I
created some simulated data meant to reproduce their magnetic tweezers data and analyzed itwith their t-test-based step-finding algorithm (101). Their data for RecQ unwinding was taken at
200 Hz with 1 mM ATP, and the standard deviation of their thermal noise appeared to be ~5 bp(putting their expected S/N close to 1). I then generated simulated RecQ stepping data with thesevalues using only exponentially distributed 1-bp steps (Figure 4.9A). I used a value of 0.0135 s
for the exponential time constant, which empirically produced a mean unwinding rate of ~60
bp/s, comparable to my previously measured 1 mM ATP RecQ speeds (section 3.4).

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Fitting this data with the step-finding algorithm produced a fit that had a roughly
Gaussian distribution of step sizes centered on 5.5 bp (Figure 4.9B). Since the real distribution ismade up of only 1-bp steps (although multiple 1-bp steps can come right after another, due to the
combination of fast stepping speed and low bandwidth), something is clearly going wrong. The
most likely explanation is that the 5-bp noise completely dominates the real 1-bp signal, andsince the algorithm is designed to fit only statistically significant differences in extension, it iseffectively fitting the noise. This conclusion, combined with the previous absence of 5-bp kinetic
step size data on my end, indicates that this kinetic step size is likely not real, but merely aproduct of fitting low S/N data. Given that this kinetic step size and the prior multi-bp step size
distribution were both postulated to occur due to RecQ occluding multiple unwound nucleotides,
and that now I have given evidence that both conclusions were drawn from over-interpreting low
resolution data, I think we can conclude that there is currently no evidence for RecQ to possess
any nucleotide-occluding capacity. It simply takes very fast 1-bp steps.

130

Figure 4.9: Fitting simulated magnetic tweezer steps.
A) Example trace for simulated 200 Hz 1-bp steps with 5-bp noise (grey) from a magnetic
tweezers assay. The true steps (green) followed a single exponential time distribution with atime constant of 0.0135 s (equivalent to a ~60 bp/s unwinding speed to simulate 1 mM ATP
conditions). The fit steps (red) arise from a t-test-based step-finding algorithm that looks for
significant differences in extension noise. B) Histogram for the fit step sizes (N = 1102) for asimulated 100 s trace. From the Gaussian fit giving a mean step size of 5.5 bp, it would seem thefitting algorithm primarily fit the 5 bp noise in the trace.

131
4.2

Unwinding “popping” behavior

In previous sections, I worked to optimize the ATP concentration so that RecQΔH would unwind
slowly but with adequate processivity for a stepping assay. During that search, I checked to see
how 300 pM RecQΔH interacted with activated 50% GC DNA hairpins when no ATP was insolution. I expected to see flat traces with no activity, since RecQ’s translocase and helicase
activities are known to be ATP dependent. However, that is not what I saw. While no ATP-lesstranslocase activity was observed (my assay cannot detect translocation), I did observe ATP-lesshelicase activity at the activated hairpin. Unlike the smoothly sloped and processive extension
increases from ATP-dependent unwinding, this ATP-less unwinding produces “popping” events
where the extension sharply increases and decreases repeatedly (Figure 4.10A).
These transitions appear near-instantaneous in the 1 kHz extension data bandwidth, amarked difference from the usual RecQ unwinding signal. However, extension data is inherently
force dependent, so judging the exact sharpness of large transitions can be hard since dramatic
shifts in force skew the signal before the force clamp can react. Polymer folding studies remedy
this by converting extension into contour length, a force-independent quantity. To create acontour length trace, the measured force and extension traces are combined in an eWLC (seesection 1.2). The benefit of this approach is that the contour length shows the real sharpness oftransitions; the cost is an increase in apparent noise.
Noise increases because the recorded force data used in the contour length conversion is
not the exact force on the DNA. We record force by measuring the trap’s force on the bead; themean value of this force is the same as the force on the DNA, but the thermal force fluctuations
are anti-correlated. For example, thermal bead kicks towards the DNA reduce the DNA’s tension
since the bead is not pulling as hard. In contrast, the trap’s force on the bead increases because

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Figure 4.10: ATP-less RecQ popping of DNA hairpins.
A) Comparison of two 10 pN RecQ unwinding traces, one with saturating ATP (red) and one
with no ATP (purple). Light: 1000 Hz, Dark: 100 Hz. Unexpectedly, RecQ displays a strange
unwinding behavior when no ATP is in solution. B) A 1000 Hz contour length trace shows the“popping” event happens instantaneously, indicating that an unwinding mechanism distinct
from normal, ATP-utilizing unwinding must be involved.

133
the bead is now further away from trap center. Combining the bead’s force and tether’s extension
to produce contour length amplifies the apparent thermal noise by effectively counting it twice,once for each trace. However, so long as the transitions of interest are larger than the noise, thisis only a minor problem. A slightly deeper problem is that extension traces can only be converted
into a single polymer’s contour length; you have to pick one polymer model for the calculation.
Since my construct is dominated by the extension of dsDNA handles, I convert to dsDNA
contour length. However, my signal arises from ssDNA unwound from the hairpin stem, so
modeling it as dsDNA introduces some error. That said, I only care about obtaining a clear
transition time and a relatively accurate length change, so the expected ~15% contour length
magnification from treating ssDNA like dsDNA at 8 pN is acceptable.
The contour length trace reveals that the “popping” transitions are indeed very fast,
unwinding 7 nm almost instantaneously for a 1000 Hz bandwidth. This gives a lower estimate of~7000 nm/s for the unwinding speed (Figure 4.10B), two orders of magnitude faster than themean unwinding speed at saturating [ATP] found in the last chapter, where RecQΔH unwound
50% GC DNA at ~45 nm/s (extension unwinding; section 3.4). From this dramatic difference,popping and normal, ATP-dependent unwinding likely occur via different mechanisms. Whatthen is the mechanism, and is it biologically relevant to RecQ? ATP-less unwinding activity is
known to be relevant to many other helicases; specifically, it is well-established for the largest
SF2 helicase family: the DEAD-box RNA helicases (69, 135, 136). The unwinding mechanism
for these proteins appears to be duplex destabilization via energetically-favorable binding, which
unwinds 8-20 bp segments of dsRNA and DNA-RNA hybrids (but not dsDNA). This
destabilization occurs without ATP, but binding ATP increases substrate affinity andhydrolyzing ATP appears necessary for unbinding and restarting the unwinding cycle.

134
In contrast to this, I observe completely ATP-less binding, unwinding, and (apparent)
unbinding by RecQΔH on DNA hairpin substrates under tension. I say apparent unbinding
because my instrument cannot definitively detect binding and unbinding, unlike more complex
optical traps integrated with single-molecule fluorescence (137, 138). I can only infer binding
and unbinding events by pauses of inactivity before and after unwinding events, respectively. For
example, performing this ATP-less assay at 8 pN produces traces that are largely blank (aside
from instrument drift) with an occasional popping event (Figure 4.11A, blue). Due to the large
time intervals between the events, we assume they are from separate helicases, and that binding
and unbinding events flank them. However, we cannot specify the exact times. One might expectthat they occur directly before and after the unwinding event, but increasing the tension to 10 pN
disproves that notion (Figure 4.11B, green). At this force, multiple popping events take place insuccession, spanning time intervals from 3 s to 8 s. Thus at lower force it is likely that the RecQ
stays bound to the substrate for much longer than the popping event implies.
Interestingly, though, the extended series of popping events are still flanked by long timeintervals of inactivity. If we assume that each condensed series of pops corresponds to one
RecQΔH binding event, then the overall rate of RecQΔH binding appears similar at 8 pN and 10
pN. This is expected, since the [RecQΔH] is the same and RecQΔH’s on-rate has no clear force
dependence. In contrast, the popping frequency within one of these bound intervals increases
dramatically with higher force. Thus the traces gives us a preliminary notion for how RecQΔH
interacts with the substrate: it binds near the hairpin junction at a rate independent of force, andonce there periodically “pops” open the hairpin with a force-dependent rate (higher force formore events). Note that I specify the ss-dsDNA junction here, since only junction unwinding canbe detected in this assay (internal melting further up the hairpin stem would not add extension

135

Figure 4.11: Quantifying the popping phenomenon.
A) Varying the tension affects the likelihood of popping in some capacity. At 10 pN (green), the
RecQΔH repeatedly pops open the hairpin stem in bursts, compared to the less active 8 pN trace(blue). However, the overall likelihood of there being some number of popping events per unit
time does not seem to shift in these traces, indicating that force many increase the number ofpopping events, but RecQ binding determines if any take place. B) A close-up example of oneof the repeated popping events. The opening size appears consistent. C) A distribution of 10 pN
opening sizes (N = 127) fit with a Gaussian. D) A distribution of the opening times, i.e. thelength of time any popping event lasts (N = 127) fit with a single exponential.

136
into the tether). Force dependence implies the popping is resisted by duplex stability, since using
force to lower that stability increases the popping rate. The opening distances are also fairly
consistent, and therefore may hold information about the underlying mechanism (Figure 4.11B).
Using 10 pN data to collect enough popping events, I measured the extension change foreach clear opening. Plotting these values in a histogram (N = 127), we find that they can be fit by
a Gaussian with a mean and standard deviation of 6.2 ± 0.6 nm (Figure 4.11C). If we assume theextension change is solely due to ssDNA being released into the tether (and not from distortions
to the ssDNA landing site), then we can convert into unwound nucleotides. At 10 pN, unwinding
1 bp generates 0.826 nm of extension change, which means a 6.2 nm change must arise from
~7.5 bp, or 15 nt of ssDNA. The length of time each pop lasts can also be quantified; fitting this
distribution with a single exponential produces a time constant of 0.19 ± 0.02 s (Figure 4.11D).
This time distribution indicates that the popping interaction is somewhat stable, as the times lastfrom milliseconds to seconds (as opposed to micro- or nanoseconds). Multiple openings also tellus that the interaction can repeatedly occur and be recovered from in a single RecQ binding
event. This likely means that some part of the RecQ is continuously, stably bound during theentire popping process.
However, it is unclear whether monomers or multimers of RecQΔH are the active agents.
The work discussed in section 3.3 indicates that monomers should be the only species present,
and current data implies that, as well. While collecting very low ATP unwinding traces for my 1bp RecQ step search, such as 5 μM ATP in a 12 pN force clamp, I would sometimes observeboth types of unwinding, sharp pops and processive unwinding, in a single trace (Figure 4.12A).
The rapid transitions from popping to processive unwinding imply that a single type of RecQΔH
generates both, assumedly a monomer given my past findings. It also implies that the binding

137

Figure 4.12: Structural implications of the popping behavior.
A) At low [ATP], normal unwinding and popping can both occur. High trap stiffness was
activated once processive unwinding began. B) The crystal structure of the RecQ homolog from
C. Sakazakii bound to forked DNA (PDB 4TMU), which we assume is close to how E. coli
RecQ binds its substrate. C) Popping can also take place in 1 mM ATPγS, with noticeably
longer opening times and occasional secondary pops. D) A cartoon of one possible mechanism
for popping. The RecQ may bind the hairpin stem in such a way that some number of base pairs
becomes destabilized. If that happens, the tension across the remaining base pairs may beenough to unzip them. Openings would last until RecQ changes its position and allows thehairpin to reclose.

138
mode for popping cannot differ too much from normal binding, which we assume is akin to theC. sakazakii homolog structure (Figure 4.12B). To gain a better understanding of what exactly
causes the pops, we can next ask whether slight structural differences change the behavior, suchas those between free RecQΔH and RecQΔH bound to ATPγS. Does popping occur when anucleotide is bound between the RecA-like domains? Putting 1 mM ATPγS in solution with
RecQΔH does produce popping behavior, but with some clear differences from the 0 ATP case
(Figure 4.12C). Qualitatively, it seems the ATPγS increases the stability of the popped structure,as those states last for much longer here (10s of seconds). Interestingly, there are sometimes
further, brief extension increases on top of the initial, longer-lasting pops. A few instances of thisbehavior were observed in 0 ATP traces, but it occurs more frequently in the ATPγS data.
From all of this disparate information, can we synthesize a plausible mechanism? The
extension data indicate that ~7.5 bp of hairpin stem, or 15 nt of ssDNA, is being unwound andreleased into the tether. The time distribution indicates that the popped state is somewhat stable,
and the low ATP traces implicate a monomer as the phenomenon’s source. Importantly, popping
is a force-dependent process. One possible mechanism is that RecQΔH binds near the ss-dsDNA
junction, but not in its assumed normal position, and slightly destabilizes part of the hairpin stem
(Figure 4.12D). Why does this work? Hairpins can be unzipped by force, with prior work
showing that 8 pN can unzip 8 bp stems (20). The pops are consistent with an ~8 bp hairpin
opening, which can be done purely through force in these 8-12 pN force-clamp assays. Given our
40 bp hairpin substrate, this could only occur if RecQ destabilized part of the stem such that the
only resistance to unzipping at the junction was from less than 8 bp of dsDNA. Internallydestabilizing the dsDNA, like the DEAD-box RNA helicases mentioned above, would thenrelease the nucleotides between the junction and destabilization point into the tether. If the

139
destabilizing binding between the RecQΔH and DNA is strong, the RecQΔH can also trap theDNA, preventing it from rezipping until the RecQΔH moves away. This would lead to thesomewhat long-lasting observed popping events. Importantly, the data indicates that RecQΔH
cannot be in its assumedly normal, junction-binding position while popping occurs. If it was, itwould act like a clamp between the ssDNA landing site and part of the hairpin stem, since it is
attached to both (Figure 4.12B). In the case of a pop, this would prevent half of the released
nucleotides from entering the tether and increasing the extension. Thus to get ~15 nt released
into the tether, 30 bp would have to be unwound, something sterically impossible here.
While this is one possible mechanism, it is obviously not the only possible one; to further
support or refute any model, we need more data. One avenue is the biochemical variations
discussed previously (section 4.1.3), applied to this behavior. Another is to use RecQWT and
RecQ mutants to try and identify which domains are important for the popping interaction. A
promising candidate would be RecQΔWH, a RecQ variant with both its HRDC and winged-helix
domains deleted; given the assumed flexibility of the WH domain, it is likely to be involved inmediating the popping. Yet even if a mechanism is determined, it remains to be seen whether thisbehavior is biologically relevant. Though popping only takes place regularly under strain, there
are several possible strained structures, such as improperly annealed strands or recombinationintermediates, where this behavior may aid RecQ in its fundamental repair and recombination
unwinding activity.
As a final note, one may ask why this behavior was never reported in prior studies. The
reason is the same as for RecQ back steps: the two kinds of prior studies were bulk biochemicaland single-molecule magnetic tweezers studies. As we have seen, the opening distance is ~8 bp,
and its frequency is force-dependent. Without external force or exotic substrates, it is likely that140
the opening rate is practically zero, and therefore bulk studies would never detect it. For
magnetic tweezers, experiments reported thus far have typically used only high [ATP], and sowould not observe the low-to-zero ATP popping behavior.

4.3

Additional non-standard interactions of RecQ and DNA

In the last section, we saw that RecQΔH could generate large unwinding “pops” when there wasno ATP in solution. Those were not the only rapid extension shifts I observed. In the presence ofATP, right before normal unwinding on activated hairpin constructs, I would sometimes observe
sharp, brief extension decreases (Figure 4.13A). Even more rarely, I would occasionally see
these spikes in isolation, with no unwinding following them. To my knowledge, this behavior
has never been reported in a helicase before. So like in the last section, I want to determine themechanism behind these strange signals, and whether it is biologically relevant.
To get a general idea of the mechanism, we need to note two things. First, these spikes
occur as extension drops below the baseline, which in my activated hairpin substrates are only
possible if part of the tether gets scrunched together. We can quantify these spikes by taking an 8
pN sample (N = 23) and finding its mean and standard deviation (Figure 4.13B). From this, it isclear the spikes cluster around a mean value of 3.5 ± 0.7 nm (not in a Gaussian distribution, atleast at this sample size). The second thing to note is the timing: the extension drops either occur
in isolation or before unwinding begins, never during or after. Therefore, they likely areassociated with a kinetic step that precedes unwinding, namely RecQΔH binding and/or
unwinding initiation. The fact that unwinding can almost immediately follow these events
indicates that they likely take place close to the hairpin junction. From these points, we can inferthat during binding and/or initiation, RecQΔH has a chance of coming in contact with spatially

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Figure 4.13: RecQ can induce sharp extension drops prior to unwinding.
A) A normal RecQ unwinding trace at 1 mM ATP. Before the start of processive unwinding,several sudden extension drops can be seen. These drops are bizarre, as there is no expected way
to generate extension drops in a fully wound hairpin substrate. They can be occasionally seen
before unwinding or in isolation, but never after unwinding. B) A small distribution of sizes forthe extension drops (N = 23) at 8 pN. They seem to cluster around a mean value of ~3.5 nm. C)
Diagram of the 40 bp hairpin construct with expected extensions at 8 pN tension. To generate
the extension drops, some part(s) of the orange labeled lengths must decrease.

142
separate parts of the tether and briefly bending them closer together. Given that the drops are rare
and have moderate dwell times (0.07 s ± 0.04 s, N = 23), we can assume that they are due to nonstandard interactions between the helicase and DNA. If this is true, looking at the substrate
structure may help identify exactly where and how these events take place.
Given that the activated substrate is made up of ssDNA and dsDNA sections, one may
ask whether one type is more easily bent than the other. To answer that, recall that ssDNA has amuch lower persistence length than dsDNA (0.75 nm vs. 50 nm; see section 1.2), meaning it ismuch easier to bend. Looking at a diagram of the substrate, there are two ssDNA locations where
the extension might be compressed, the 6 nt left side and the 33 nt right side (Figure 4.13C). At 8
pN tension, these sections will be 2.3 nm and 12.4 nm long, respectively. In contrast, it is hard to
imagine the RecQ changing the extension of the dsDNA handles or the 2 nm width of the hairpin
stem. So what interactions could generate these spikes? The maximum extension drop one could
get from the left ssDNA section is ~2 nm, assuming the hairpin stem touches the left dsDNA
handle and creates a tensionless 6 nt loop. This is not enough to account for the recorded spike
sizes. Therefore, the majority of spikes must involve at least the 33 nt side (possibly both sides).
Considering that RecQ’s RecA-like domains bind to ssDNA and its zinc-binding andwinged-helix domains associate with dsDNA, there are two probable geometries for these spikes.
Each involves RecQ binding non-neighboring parts of the DNA substrate, likely due to strong
thermal kicks briefly bringing these parts close enough together to both interact with RecQ. The
first solely involves the RecA-like domains binding to non-consecutive nucleotides, forming aloop of tensionless ssDNA. The second involves the winged-helix domain; if it can rotate up to
90° like in the homolog structure (83), it could bind faraway dsDNA such as the left handle ordistant parts of the hairpin, contracting the tether extension and leading to the observed drops. A

143
simple way to discover if this geometry is right would be to use RecQΔWH, the RecQ mutant withboth its HDRC and winged-helix domains deleted, and see if the spikes persist. Sadly, as I could
not obtain these enzymes myself, I have not performed this experiment.
Another strange interaction between RecQ and its DNA substrate often occurred justbefore the end of hairpin unwinding: a pause followed by the sudden unzipping of the rest of thehairpin (Figure 4.14A). Unlike the spikes, this behavior is rather common; for instance, it can beseen in multiple previous plots in this thesis (c.g., Figures 4.4 and 4.6). In addition, it has also
been observed by an independent group using a completely different DNA hairpin and type of
RecQ (Figure 3b in Ref. (134), using a plant RecQ from A. thaliana). Clearly, force-induced
unzipping of the remaining short hairpin stem causes the sudden opening, as discussed in section
4.2; that in itself is not surprising. The surprising part is that these events are preceded by notable
pauses, even at saturating [ATP] where pauses are nearly non-existent. Why? If the RecQ simply
unwound the hairpin stem until it could no longer resist the tension, one would expect a pop to
occur instantaneously. Instead there are significant dwells in the traces. Quantifying the sizes ofthe pops and dwells may help elucidate these events.
Given that helicase pausing and unwinding distances can typically both be modulated byapplied force, I collected top extension changes (Figure 4.14B) and stall times (Figure 4.14C)
from both 8 pN (red, N = 41) and 10 pN (grey, N = 52) data sets using 1 mM ATP. The
distributions for the top pops could both be well-fit by Gaussians: the 8 pN distribution has aGaussian fit of 6.1 ± 1.7 nm (mean ± s.d.), while the 10 pN one has a fit of 7.0 ± 1.3 nm (mean ±s.d.). While these means are within one standard deviation, the overall distributions are clearly
distinct from one another. To further investigate whether they could be related, I tried converting
the extension changes into unwound base pairs. This conversion had to account for the difference

144

Figure 4.14: RecQ can stall force-induced hairpin unzipping.
A) During processive unwinding, RecQΔH will often stall before the hairpin completely unzips,
followed by a sharp unzipping event. The extension change between the stall position and thecompletely unwound hairpin is relatively consistent. B) Measuring extension pops after stalls at
8 pN (pink, N = 41) and 10 pN (grey, N = 52). Both distributions produce a similar mean value
when converted into force-independent base pairs. C) The dwell time distributions for the stalls
from (B). They appear to have a roughly exponential decay. D) If E. coli RecQ has a similarbinding structure as the C. Sakazakii homolog, its translocation might stall when its dsDNAbinding domains encounter the hairpin loop. After a time, the applied force may then open theremaining hairpin stem.

145
in 8 pN vs. 10 pN extension, as well as the extension change brought about by unwinding the lastbase pair in the hairpin (2 nt from the base pair + 4 nt from the loop are added; 2 nm from thefixed stem width are subtracted). Interestingly, unwinding the last 9 bp of the hairpin at 8 pN
results in a total extension change of 6.25 nm, very close to the 6.1 nm fit mean. Moreover,unwinding the last 9 bp at 10 pN results in an extension change of 7.1 nm, also very close to the
7.0 nm fit mean. Therefore, both distributions could be interpreted as being the same distribution,with the extension values shifted due to applied force. This would implicate a force-independent
mechanism somehow related to the 9 bp value of both means.
The dwell time distributions for the two forces are more similar than the corresponding
extension ones, although they do not have a well-defined shape. Looking at the bulk of the data,
most dwells appear to last 25 to 100 ms, which is a non-trivial amount of time on the singlemolecule level. To instigate such dwells, the cause must present a significant barrier to normal
unwinding, and must be able to do so repeatedly given the frequency of the events. On that note,the above data sets were collected with the intent of investigating the pause-then-pop
phenomenon; therefore, traces not exhibiting that behavior were excluded. While common, thisphenomena was by no means universal; many traces unwound the entire hairpin without any
pause or pop. Whether or not this was a helicase-dependent effect – that traces either had top
popping or they did not – has not been quantitatively analyzed. However, my qualitative answer
is no: unwinding runs from a single molecule could both complete the hairpin without incident or
undergo a pause-then-pop.
Given this information, what conclusions can we draw? The relative force-independenceof both the dwell time and pop size distributions suggests that the substrate parts under tension,
the dsDNA handles and the ssDNA sections, are not the source of the pauses. Since the source

146must involve RecQ interactions with DNA in some way, that leaves only the hairpin itself as apotential cause. And this leads us to revisit an interesting fact about the RecQ homolog structurefrom Ref. (83). When the homolog binds to the forked substrate, it unwinds 2 bp of the original
10 bp hairpin stem, leaving 8 bp associated with the enzyme (Figure 4.14D). This conformation
likely arises from packing constraints in the protein crystal; in other words, this amount of
hairpin stem somehow fits well with the protein, perhaps by limiting the stem length jutting out
from the bulk of the protein. This would put the hairpin loop right next to the RecQ domains.
Could the hairpin loop serve as an obstacle to further unwinding, perhaps because the geometryof the loop differs so much from the double helix? This is my current imagined model for how
these top popping events take place, given the striking similarity of the crystal’s 8 bp stem length
to the current assumed 9 bp stem unwinding. RecQ unwinds processively until its dsDNAassociating domains encounter the hairpin loop, whose geometry makes it more difficult toassociate with. Depending on thermal fluctuations, the helicase can either stall here or overcome
the kinetic barrier and continue unwinding. If it stalls, the tension across the tether is sufficient toeventually unzip the remaining hairpin, allowing RecQ to continue translocation.
Obviously this is a speculative model; my current data does not allow me to determine amore definitive one. Like the other strange phenomena discussed in this and the prior section,collecting data with RecQWT and RecQΔWH would be very informative. The addition or loss of a
dsDNA-associating domain could dramatically affect this behavior, especially if it is indeed dueto helicase domains being blocked by the hairpin loop.

147

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155

Appendix A
Acronym / Abbreviation Index%GC

percent guanine-cytosine content

1D, 2D, 3D

one, two, or three dimensionalA

adenineACalternating current

ADC

analog-to-digital converter

ADP

adenosine diphosphate

AFM

atomic force microscopy

AOM

acousto-optic modulator

ASCII

American standard code for information interchangeATPadenosine triphosphateATPγSadenosine 5′-O-(3-thiotriphosphate)BME

𝛽-mercaptoethanol; 2-mercaptoethanol

BNC

Bayonet Neill–Concelman connectorbpbase pair

BSA

bovine serum albuminCcytosine

156
CAD

computer-assisted design

CCDcharge-coupled device

DAC

digital-to-analog converter

DAQ

data acquisition

DBCO

dibenzocyclooctyl

DCdirect currentDNA

deoxyribonucleic aciddsDNA

double-stranded DNA

DTTdithiothreitol

EDTA

ethylenediaminetetraacetic acidF-x

force-extension

FFT

fast Fourier transform

FJC

freely-jointed chain (e prefix: extensible)

FPGA

field-programmable gate array

FRET

Förster resonance energy transfer

G

guanineGPIB

general purpose interface bus

GTPguanidine triphosphate

HRDC

helicase-and-RNaseD–C terminal domain

HSQhydrogen silsesquioxane

KOH

potassium hydroxide

LV

LabVIEWMAX

Measurement & Automation Explorer

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MTWB

molecular tweezer wash bufferMWmolecular weightNAnumerical aperture

NI

National Instruments

NIH

National Institutes of Health

NIM

nuclear instrumentation module

NPGS

Nanometer Pattern Generation System

NSF

National Science Foundationnt

nucleotideOS

overstretching

OT

optical trapping, optical tweezers

PBS

phosphate buffered saline

PBS

polarizing beam splitter

PCA

Protocatechuic acid (3,4 – Dihydroxybenzoic acid)

PCD

P8279 – Protocatechuate 3,4 – Dioxygenase from Pseudomonas sp.

PCI

peripheral component interconnect bus

PCR

polymerase chain reaction

PDB

Protein Database

PEG

polyethylene glycol

PIN

p-type, intrinsic, n-type semiconductor

PIO

parallel input-output

PXI

PCI eXtensions for Instrumentation

PZT

lead zirconate titanate piezo-electric transducer

158
QPD

quadrant photodiodeRFradio frequency

RNAribonucleic acidSA

streptavidinSDstandard deviation

SDS-PAGE

sodium dodecyl sulfate–polyacrylamide gel electrophoresis

SEM

scanning electron microscopeSFsuperfamily

SMFS

single-molecule force spectroscopy

S/Nsignal-to-noise ratio

SSB

single-stranded DNA binding protein

ssDNA

single-stranded DNA

T

thymine

TEM00

transverse electromagnetic mode (0,0)

TEV

tobacco etch virus (a source for the associated protease)

TMAH

tetramethylammonium hydroxide

UTPuridine triphosphateUV

ultravioletVIvirtual instrument

WLC

worm-like chain (e prefix: extensible)

159

Appendix B
Protocols Index

In this section, I provide some of the common protocols I utilized during my thesis work. They
are reproduced in their original formatting for ease-of-use. The components for the bolded buffer
solutions used in many are given below.

DNA Dil:

20 mM HEPES pH 7.5, 20 mM KCl, 1 mM Na+-EDTA pH 8

MES-Tw:

100 mM MES pH 5, 0.3% Tween-20

MTWB-150:

20 mM Tris-HCL pH 7.5, 150 mM NaCl, 0.4% Tween-20

Phos:

81 mM Na2HPO4, 19 mM NaH2PO4

Phos-Tw:

Phos with 0.3% Tween-20

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Streptavidin to CML beads: Coupling Procedure
CML: Carboxylate Modified Latex(Based on protocol in Section 14.4.3 of Bioconjugate Techniques, 2 nd Ed.)

Buffers:

MES-Tw

Phos

Phos-Tw

Reagents:

Sulfo-NHS

EDC

BME

1 M Glycine

Notes: Make Sulfo-NHS and EDC solutions fresh every time. Easiest to make two batches at a time. This
procedure will take at least 7 hours to complete: schedule day appropriately.

Procedure
Prep Beads and SA
1. Take out a new 1 mg bottle of Streptaviden (Vector Lab), so that it can warm up while you work.
2. To a 1.5 mL microfuge tube, add 100-200 L CML latex beads (~4.0%) and 150 L MES-Tw buffer (200 L for
750 nm beads; 100 L for 500 nm beads).

Wash and Sonicate Beads
3. Pipette mix and spin at 14K for 4 min. Take supernatant out, and resuspend in 200 L MES-Tw.
4. Repeat Step 2 two more times, for a total of 3 washes.
5. After resuspending in 200 L MES-Tw, sonicate the beads (sonicate until you complete Steps 6 and 7).

Make Special Solutions
6. Make 0.1 M sulpho-NHS sol. (21 mg/mL). Dissolve 4.2 mg of NHS (in 4 C fridge) in 200 L MES-Tw.
Note: If you measure out more than 4.2 mg, use the ratio 21 mg / 1000 L = M mg / V L to find theneeded suspension volume V. Do not measure out less than 4.2 mg (for at least 200 L).
7. Make EDC sol. (5 mg/100L). Dissolve 10 mg EDC (-20 C fridge) in 200 L MES-Tw.

Add Special Solutions and Mix
8. End sonication. Add 700 L MES-Tw to the bead solution, for a total volume of 900 L.
9. Add 100 L of sulpho-NHS solution to the 900 L bead solution. Briefly vortex.
10. Add 100 L of EDC to the 1000 L bead/ NHS solution. Briefly vortex (Total volume = 1100 L).
11. Put on rotator for 30-45 min at room temperature. Do not exceed 45 minutes.

Quench EDC with BME
12. After rotating, add 1 L -mercaptoethanol (BME) to the bead solution (under the hood). Briefly vortex androtate for 10 min at room temp (parafilm the tube first). (This quenches the rxn of EDC).

Make SA Solution and Wash Beads Again
13. While the bead sol. rotates, make streptavidin (SA) sol. by adding 1 mL Phos to SA bottle. Mix on rotator for
20 min. After rotating, transfer the SA solution to a new 1.5 mL microfuge tube. (Total SA volume = 1000 L).

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14. After the bead solution rotates for 10 min, spin down for at least 5 min at 14K. Pull off as much of thesupernatant as possible without taking a significant amount of beads.
15. Resuspend beads in 200 L Phos-Tw, and do three 3 minute washes with 200 L Phos-Tw.
16. For final resuspension, bring beads up in 500 L Phos-Tw.
17. Optional Step: Add 1 L beads to 100 L Phos-Tw and flow 20 L of this solution through flow cell. Checkfor clumping under a microscope. If clumped, sonicate 8 min or until unclumped.

Add SA Solution and Mix
18. Add beads to SA sol. and quickly mix with pipette (need to monodisperse quickly). Vortex briefly.
(Total SA/bead volume = 1500 L).
19. Place on rotator for at least 2 hours at room temperature. 2 ½ hours is ideal, 4 hours is too much. If desperate
and pressed for time, leave in 4 deg C fridge’s rotator overnight.

Quench SA-CML Reaction
20. Quench reaction with glycine by adding 30 L 1 M glycine (MW=75 g/mol). This step is critical.
21. Vortex and let sit on rotator for 10 min at room temperature.

Spin Down and Wash Final Product22. Spin down sol. for at least 15 minutes at 14K. Extract supernatant and resuspend in 200 L Phos-Tw.
23. Wash 5 times with Phos-Tw. Change tubes after 3 washes.
24. For final resuspension bring up in 200 L Phos-Tw (If doing multiple batches, combine all finished batches
into one tube). Label final tube and store in 4 deg C fridge.

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Cleaning Coverslips
1. Put large stir bar into 1 L washed beaker, zero out on scale, and weigh out 50 g KOH pellets into it (+/- 1 g). Fillbeaker with 170 mL EtOH and 80 mL dH2O. Cover with aluminum foil. Label with contents, initials, and date.
2. Put on the stirrer/hot plate in the fume hood. Mix at 200 rpm until you’re done preparing the other components.
3. Load coverslip basket with Corning coverslips, and attach handle (we typically do 3 at a time).
4. Place two 1 L washed beakers ½ full of dH2O in Branson 5200 sonicating waterbath, as well as KOH solution
beaker and a 1 L beaker with ~ 300 mL acetone.
5. Fill water bath 1/4 full with water from sink. Turn on sonicator (but no heat).
6. Using coverslip basket, place coverslips in acetone for 3 min.
7. Rinse over sink with ethanol (remember: alcohol washes alcohol).
8. Place coverslips in KOH solution for 3 min.
9. Rinse coverslips over sink with dH2O squirt bottle.
10. Place coverslips in beaker of dH2O in waterbath for 3 minutes. Rinse over sink with dH2O. Place in other beakerof dH2O for 3 min. Rinse over sink with dH2O again.
11. Rinse over sink with ethyl alcohol. Dry in microwave. High power, 2 min.
12. Wash coverslip container with ethanol, remove handle, place coverslips in, and cover with parafilm. Write yourinitials, the date, and “KOH cleaned” on lid.
13. Repeat steps 6-12 for the rest of the coverslips, changing the ddH 2O in the beakers in the waterbath every threebaskets.*
14. Clean-up. Dispose of:dH2O in sink
Acetone in organic hazardous waste (jar is labeled, use funnel)
Solution of KOH and alcohol in organic hazardous waste (jar is labeled, use funnel)
Empty waterbath in sink (use the plug in its back or just pour out)
Put empty beakers, etc. in “needs cleaning” tray
*It’s most efficient to parallel process three baskets at a time. Once the first basket is in the KOH, wait no less than
1:30 (less than that and you will hate life as it takes about 90 seconds to get all of the KOH rinsed off the coverslips
with a squirt bottle) and start the next basket in acetone.

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Making HSQ Post Coverslips
Most important steps: (1) spin coat process [amount of HSQ and introduction speed], (2) correct picoammeterreadings, and (3) development process.

1 Preparation for lithography
1. Use diamond scribe to mark lower right corner of each coverslip with “P” to indicate post side.
2. KOH clean coverslips.
3. Gather materials needed to spin coat. Bring into clean room if not already there.
a. HSQ (FOx16). Acquire from 4 C fridge in B1B12. Take 210 uL/slide in microfuge tubes. Return
HSQ bottle to fridge as quickly as possible.
b. Coverslips in Teflon rack in box.
c. 200 uL pipette.
d. Clean room box: contains 200 uL tips, tweezers, and SEM mount.
e. Diamond scribe from SEM room.
4. Turn on the spin coater bench’s power and activate its nitrogen ventilation. Turn on the spin coater itself
and the hot plate. Heat to 180 C. Place faux-paper towel in front of hot plate.
5. Find usable spin chuck for coverslips and insert into spinner (O-ring on chuck needs to be smaller thancoverslip width, so that vacuum suction can hold slip in place).
6. Set spin coater acceleration to 3000 rpm/s, velocity to 2000 rpm, and spin time to 45 s. On newer spincoater, these values are (sometimes) saved in Program 20. On older one, manually turn the dials until theelectronic displays show the right values.
7. Place coverslip on chuck with tweezers, and load pipette with 200 uL HSQ. For new model, go into Run
Mode, press Vacuum, then Start to start spin. On old model, use foot pedal to start spin.
8. Immediately dispense HSQ onto middle of slip. After 45 s spin, disengage vacuum and remove slip withtweezers.
9. Heat coverslip (dry side down) for 4 min at 180 C.
10. Use diamond scribe to mark an “X” in corner beneath the “P”.
11. Lay finished coverslip on paper towel to cool, then place in rack. Repeat process as necessary.
12. Clean up area, and shut off hot plate, spinner, nitrogen, and bench.
2 Electron beam lithography
1. Vent the SEM, and open up the chamber.
2. Install the Helix LVD for LoVac mode. First, vertically slide the aperture up and in, then twist cw until
tight. Then slide the chip gently into the slot. Ask David for help if you are unsure.
3. Install mount into SEM. Loosen locking nut with hex wrench if mount pin doesn’t slide in fully. Align sothat long edge of mount is parallel to door, and the permanently depressed metal pin is nearest thecomputer. Tighten until the mount cannot be rotated by hand.
4. Close the chamber door and pump down in LoVac mode (hold door closed until door gap minimizes). Put
LVD in 2nd quadrant. If LVD is not an option, detector was not installed properly.
5. While pumping down, open NPGS and go into Matt Bull’s files. On the top bar, select ‘Commands’ andchoose ‘Calibrate DACs’.
6. Once pump is done (chamber icon is green), activate 30 kV HV and put spot size at 3-4. Check thatContrast is 60-90 and Brightness is 30-70. Unpause 2nd quadrant.
7. Use the stage panel to move to (0,0). Type values into X and Y boxes, then hit Enter.
8. At ~100x Magnification, manually move stage in XY (click on 1 st quad and hold mid mouse button) until
you find the inside edge of the mount’s metal washer.
9. Bring stage up in Z to 7-9 mm by clicking on CCD quad and using mid mouse button (can also use Z input
box, but only do so when far away from LVD; hitting Esc stops motion).
10. Magnify on washer edge to ~2000x, and change dwell time to ~0.8 µs. Adjust fine focus knob until imageis crisp (striations visible on edge, etc). This will change the listed working distance (WD). Once a good
image is obtained, link WD to z-height.
11. Zoom out to ~200x and find the mount’s center. Look for an FIB-ed “X”. Zoom to 2500x, put the quad
center cursor over it, and record the XY coordinates in Notepad.

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12. Turn off HV, vent the chamber, and open the door. Lower the loading pin and lift the central metal washer.
Slide coverslip in, ensuring that the registry corner of the slip (the one with the “P”) is in the corner withthe two fixed pins. Lower metal washer.
13. Close door and pump down in LoVac mode. Once done, navigate stage to (-10 mm, 18 mm), which should
be close to registry corner.
14. Raise the stage to ~5 mm. Activate 30 kV HV and set spot size to 1.8 (or whatever results in 0.800 – 1.100
nA on the picoammeter during the write program’s run). Unpause 2nd quad.
15. Move in XY until you find the registry corner (or something easy to focus on). Zoom to ~2000x, and see if
you can get a decent image by adjusting fine focus. If not, astigmatism could be really bad.
16. If image is okay, adjust fine focus to 5 mm WD. Raise stage until image is focused. Link WD to z-height.
17. Move off registry mark to featureless area. Zoom in by couple 1000x and look for small marks. Zoom in on
one of them, fine focus adjusting as you go for best image quality.
18. At 80000x, fine focus adjust as best you can, then switch to 4x50 ns dwell and Live image (not 4 frame
average). On 1st right-hand panel, hit Lens Align modulator button.
a. If whole image moves in XY, lens alignment is off. Move Lens box cursor until it stops.
b. If image blurs in orthogonal directions, astigmatism present. Move Stig box cursor until it stop.
19. Zoom in to ~120000x, increase dwell to ~1.5 µs, and fine focus for best image. Then use Stig knobs to
improve image as best you can.
20. On 2nd quad, hit pause button. On box left of computer screen, turn knob from “on” to “ext”. In NPGS,
click “NPGS mode” ( “external” appears on quads, indicating switch from SEM to NPGS mode). Check
that the picoammeter reads 0. Unpause CCD screen and navigate to center point using XY input boxes.
21. Highlight 2nd quad and unclick beam blank icon.
22. In bullm’s folder, select post file “7x7_15umLateral_700nm.RF6”. Right click and select “Process run
file”. Screen will go black. Check that current is now flowing. If not, hit “Esc” twice.
23. Let the program run (~15 min), then turn off HV. Switch back to SEM mode, and change beam blanker box
knob back to “on”.
24. Vent the chamber and repeat the process as necessary.
25. When finished, uninstall LVD and mount. Close door and pump down in HiVac mode.
3 Development
1. In B1B12, gather three 1 L beakers. Fill two with 500 mL filtered water each.
2. Put the beakers in the fume hood. Fill the third with MF CD-26 solution stored below the hood.
3. Get a timer, and attach a handle to the coverslip rack. Submerge the coverslips in the MF CD-26 solution
for 15 min, agitating them gently every 2-3 min.
4. Remove rack, let excess MF CD-26 drip back into beaker, and submerge into water #1 for 3 min.
5. Repeat with water beaker #2.
6. Dry coverslips with nitrogen gas gun.
7. Put MF CD-26 back into its container, and store under the hood.
4 O2 Plasma Cleaning (optional)
1. Use clean room username and pin to logon to PlasmaStar computer if David’s not logged in.
2. Press the big green button.
3. Turn hot water valve (out) 90 degrees ccw and cold water (in) 45 degrees ccw.
4. Open the O2 tank in valve MF2 by rotating counterclockwise (take care to not open all the way).
5. Open PlasmaStar program. Wait for it to set up. Login: “user” with password “rie”.
6. Allow startup to complete. Open door and place coverslips on lower surface. Close door.
7. Select “Recipe management”, then “Open recipe”. On the prompt, type in “SRO_posts”. The full recipename should auto-fill; select it. Recipe values: MF2 sccm 25, 300 W, and 30 s.
8. When prompted, select “Execute recipe”. The recipe will run, and will notify you when it’s done.
9. Remove coverslips, close door, and exit Plasmastar program (it will pump down before closing).
Turn off oxygen gas and water valves (turn cw). Press the big red button. Logout if you logged in.

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Silane-PEG-Azide Functionalization Protocol
Reagent source: PG2-AZSL-600 (200 mg).
1.

Check that enough of each solvent is available: > 500 mL toluene (50 mL isopropanol).

2.

UV-irradiate desired coverslips with ozone chamber for 30-45 min.
a. Place down large kimwipe in hood. Work over it.
b. Place coverslips in large petri dish bottom. Make sure “P” side is up.
c. Place dish in chamber, shut, and press power. When time is over, hold button to turn chamber idle.

3.

Prepare reagents for silane-PEG-azide (SPA) / toluene solution.
a. Gather equipment: Cylinders: 100 mL, 10 mL. Beakers: (4) mid-sized for toluene / iso stock andwaste. Covers: Watch glass, petri dish top. Tiny, small, and medium stir bars. Metal spatula.
i. Holder version: Get holder, glass hook, holder bottom spacer, cut 600 mL beakerb. Fill stock beakers. Keep toluene bottle in hood.
c. Condition all equipment 2x. Spatula, 10 mL cylinder, tiny stir bar get isopropanol, all else toluene.
d. Set large hot plate to 60°C solution temp. For Talboys, turn the heat knob. For Omega, press Enter(Set), 1 (Plate Temp), temperature number, then Enter.
e. Take out SPA aliquot from -20°C freezer. Thaw for ~5 min (moisture condenses if opened cold).
f. Calculate the volume needed for 0.15 mg/mL solution (V = m/0.15). Should be < 70 mL.
i. Holder version: Will need 100 mL to cover holder
g. Measure 2 mL isopropanol into 10 mL cylinder, then V-2 mL toluene into 100 mL cylinder.

4.

Prepare SPA / toluene solution.
a. Pour 2 mL isopropanol into SPA aliquot vial. Agitate with spatula to aid dissolving process.
b. Put in tiny stir bar, re-cap, and place on hot plate. Stir at 800 rpm for 2 min.
c. Pour 2 mL toluene into 10 mL cylinder, then into vial. Stir at 800 rpm, 2 min. Shake to mix better.
d. Pour vial into now emptied toluene stock beaker, as well as toluene from 100 mL cylinder. Put in
medium stir bar, cover with watch glass, and put on hot plate. Stir at 600 rpm for 5 min.
i. Holder version: Use 600 mL beaker instead of toluene stock beaker.

5.

Remove coverslips from chamber and prepare them for deposition.
a. Place petri dish on hot plate. Put small stir bar in middle. Slowly pour SPA solution into dish. Try
not to move coverslips. Cover with petri dish top. Spin at 400 rpm.
i. Holder version: Put coverslips into holder. Place Teflon spacer and small stir bar into
600 mL beaker, then place holder. Cover with watch glass, spin at 800 rpm.

6.

Let deposition proceed for 30 min.

7.

Clean and dry functionalized coverslips.
a. Gather (4) 250 mL beakers. Fill toluene into first, isopropanol into second, and deionized water
into third and fourth. Fill enough to easily dunk coverslips.
i. Holder version: Use 1 L beakers instead.
b. Use two tweezers to fish out coverslips, then dunk each in the four beakers (20 s each). Transfer
through air very quickly, as clumps can form otherwise. Put into 10-fold Teflon holder.
i. Holder version: Dunk entire holder instead of individual coverslips.
c. Tape down large kimwipe on counter and turn on N2 gas.

Dry each coverslip. Once done, cover 10-fold box with parafilm and store at 4°C.

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Tethered Bead Assay: DBCO Edition

Buffers:

DNA Dil

MTWB-150

PBS 1X

Procedure
1. Make flow cells and allow epoxy to dry. Make sure functionalized “P” side points inwards.
2. Pre-wash slides with 200 µL DNA Dil.
3. Dilute DNA in DNA Dil (~100 pM). Flow 15 L DNA per slide.
-- Let sit 30 min in humidity chamber at room temperature (up to 4 h is fine).
4. While DNA incubates, prepare 15 µL SA-coated beads per slide.
A. Briefly vortex SA bead stock.
B. To 1.5 mL tube, add 0.5 µL bead stock.
C. Add 100 µL MTWB-150. Centrifuge for 5 min at 14 krpm and remove supernatant.
D. Resuspend in 50 µL MTWB-150 and transfer to a 0.5 mL thin-wall tube.
E. Sonicate ~60 min (~two rounds of program 2 in sonicator).
5. Wash slides with 200 µL PBS 1X and 200 µL MTWB-150.
6. Flow 15 µL of bead solution into each slide.
-- Let sit 1 h at room temperature in humidity chamber.
7. Wash slides with 400 mL MTWB-150.

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RecQ Protein Purification
We obtained our RecQ plasmid (RecQ Δ HRDC-TEV- pET-15b) from Keir Neuman and Yeonee Seol at the NIH.
I Transform plasmid into Rosetta2 BL21-DE3 pLys competent cells (E. coli)
1. Thaw the Rosetta 2 competent cells on ice for 10 min.
2. Add 1-2 μL of plasmid (~5-50 ng) and mix well by flicking the microcentrifuge tube for about 10 s. Keepon ice undisturbed for 30 min.
3. Transfer the tube to 42 °C for exactly 30 second and then chill on ice for 2 min.
4. Add 500 μL of SOC recovery media and let shake at 37 °C for 1 h.
5. Spread 100 μL of this onto LB-agar-ampicillin plate and incubate at 37 °C overnight.
6. The next day, take a single colony from plate and grow up in 5 mL LB with ampicillin overnight.
7. The next day, add 2 mL of this overnight culture to 150 mL of Terrific Broth (Sambrook, CSH,
opennetware) with ampicillin and grow for about 2.5 h at 37 °C in a shaker until the OD at 600 nm is ~0.6.
8. Cool culture on ice for 5 min and grow in a shaker at 23 °C for 30 min.
9. Express protein by adding 1 M IPTG to culture (0.2 mM final concentration). Continue growing for 4 h.
10. Pellet bacteria by centrifugation at 6000 g for 30 min. Discard supernatant and store at -80 °C until readyfor purification.
II Harvest protein from bacteria (all steps performed at 4 °C)
1. Resuspend bacteria pellet in cold equilibration buffer (20 mM Tris-HCl pH 8.0, 300 mM NaCl, 10%
glycerol, 20 mM imidazole, 1 mM beta-mercaptoethanol), with Halt Portease Inhibitor Cocktail (Thermo
Scientific 87785).
2. Sonicate lysate in a Branson Digital Sonifer 450 with a 1/8” Branson Microtip 101148062 (six 30 s pulses
at 25% amplitude).
3. Centrifuge at 16000 g for 30 min to remove cellular debris.
4. Load supernatant onto two Ni-NTA spin columns (Thermo Fisher PI88224 preequilibrated in equilibrationbuffer). Mix vigorously to resuspend the Ni-NTA matrix and incubate on a rotator for 30 min.
5. Remove lysate by briefly spinning in a benchtop microcentrifuge. Repeat process until all remaining lysate
is exposed to Ni-NTA matrix.
6. Wash Ni-NTA 6 times with 2.5x column bed volume (500 μL) equilibration buffer on a vacuum manifold.
7. Elute protein from Ni-NTA by adding one bed volume (200 μL) of elution buffer (equilibration buffer with
500 mM imidazole) and briefly spinning in a benchtop microcentrifuge. The flow thru containing the RecQ
was saved. Repeat elution process twice more.
8. Measure elution protein concentrations on a nanodrop at A280. Pool peak fractions and dialyze inequilibration buffer without imidazole (uncleaved MW=61600, A280 ε = 47300). Two rounds of dialysis
were performed for 2 h at 4 °C with 300 mL.
9. A portion of the fraction was incubated with TEV-his protease (Promega) overnight at 4 °C to cleave off
the his-TEV tag.
10. Clean cleaved RecQ by adding 20 mM imidazole to solution and incubating again in a preequilibrated NiNTA spin column for 30 min. Keep RecQ-containing lysate after centrifugation. Repeat process once.
11. Dialyze the highly purified RecQ in equilibration buffer with no imidazole as before.
12. Aliquot RecQ, freeze in liquid nitrogen, and store at -80 °C. Verify RecQ size on a 10% SDS-PAGE. Look
for band at ~58 kD (cleaved MW=58459, A280 ε = 45800).