Single-Molecule Investigations of Enzyme Dynamics:
RecBCD Helicase Motion and glmS Ribozyme Cleavage
byMartha Hosotani
B.A., University of California at Berkeley, 2004
M.S., University of Colorado at Boulder, 2007

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
2011

UMI Number: 3453726

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This thesis entitled:
Single-Molecule Investigations of Enzyme Dynamics: RecBCD Helicase Motion and glmS
Ribozyme Cleavage
written by Martha Hosotani
has been approved for the Department of Physics

Thomas T. Perkins

Matthew A. Glaser

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.

Hosotani, Martha (Ph.D., Physics)
Single-Molecule Investigations of Enzyme Dynamics: RecBCD Helicase Motion and glmS Ribozyme
Cleavage
Thesis directed by Professor Thomas T. Perkins

Enzyme dynamics are a critical factor in enzyme function. Probing individual enzymes andenzymatic systems elucidates the mechanistic basis of their dynamics. Here, with two distinct
single-molecule, biophysical approaches, we examine two particular enzymatic systems.
RecBCD is a multifunctional enzyme that repairs double-stranded DNA breaks and degrades
unwanted DNA in Escherichia coli. To probe how this superfamily-1 helicase couples ATP binding
and hydrolysis to unwind and directionally translocate along double-stranded DNA, we used opticaltweezers featuring 1-base-pair resolution (∆f = 0.1–2.3 Hz, F = 6 pN) to directly measure interactions between single RecBCD helicases and their DNA substrate. Surprisingly, the RecBCD·DNA
complex exhibited large, multi-base-pair conformational dynamics (4.4 ± 0.3 bp, mean ± σ x̄ , ∆f =0.1–10 Hz) that were present both between unwinding steps and in the complete absence of ATP.
To understand the origin of the conformational dynamics, we made three observations. First, theonset of the conformational dynamics depended on 50 -ssDNA length, which interacts with the RecD
motor, and were consistent with the onset of helicase activity, as established in prior biochemical studies. Second, conformational dynamics of RecBCD bound to a cross-linked DNA constructwere suppressed, indicating that the RecBCD “pin,” which interacts with the junction, undergoes
rapid back-and-forth translocation along the DNA. Third, conformational dynamics were reducedin the presence of nonhydrolyzable ATP analogs. These results indicate that when RecD is engaged
RecBCD actively destabilizes the duplex, and that these conformational dynamics are modulatedby the nucleotide-bound state of the enzyme. These studies signify the first real-time measurements
of previously-unseen destabilization dynamics of the DNA duplex by RecBCD and provide insight
into the unwinding mechanism of superfamily-1 helicases.

iv
In the effort to develop highly-sensitive biosensors for medical diagnostics, environmental monitoring, and industrial quality control, nucleic-acid-based biosensors offer a number of competitive
advantages over antibodies for sensing applications. We explored a naturally-occurring biosensor,
the glmS ribozyme, a RNA molecule that has the remarkable ability to recognize a specific metabolite, catalyze self-cleavage upon its binding, and regulate gene expression. We developed a new,single-molecule assay to detect individual glmS ribozyme cleavage events with video microscopythat can be generalized to other nucleic-acid-based biosensors.

Dedication

I would like to dedicate this dissertation to my sister, my mother, and my father.

Acknowledgements

I would like to thank my Thesis Advisor, Professor Thomas T. Perkins, without whom thiswork would not have been possible. I greatly appreciate his many contributions, which have been
instrumental in the single-molecule biophysics projects presented in this dissertation. His knowledge
has been of great resource throughout the years during which I have had the opportunity to workwith him.
I would also like to thank the many people with whom I have had the opportunity of working directly and indirectly, for sharing experimental techniques and methods, partaking in helpful
scientific discussion, and contributing to a positive research environment. I would like to thankCollaborators: Hung-Wen Li, Hsiu-Fang (Bendy) Fan, Christopher J. Wilds, Anne M. Noronha,
and Robert T. Batey; Postdoctoral Researchers: Yeonee Seol, Gavin M. King, D. Hern Paik, Rebecca K. Montange, and Ruby M. A. Sullan; Graduate Students: Amanda E. Carpenter, Ashley
R. Carter, Justin A. Kuczynski, and Allison B. Churnside; Professional Research Assistant: Wayne
A. Halsey; and Undergraduate Students: Andrew Leifer, Louisa Eberle, George A. Emanuel, Duc
Minh Nguyen, and Matthew S. Bull. I would like to thank Hung-Wen and Bendy, for characterizing
and providing biotinylated RecBCD enzymes; Chris and Anne, for synthesizing the cross-linked
DNA constructs; Rob, for training and guidance on the glmS ribozyme project; Ashley, for introducing the RecBCD optical-trapping assay and analysis methods to me, making time not only
to work side-by-side on the RecBCD project through numerous late nights, but also to engage
in invigorating, memorable experiences together outside of the lab; Amanda, for training me on
trapping instrumentation, programming, and surface-chemistry technique, helping me with innu-

vii
merable scientific details, and for sharing her perceptive and sincere insight in both lighthearted andthought-provoking interactions; Wayne, for teaching his excellent biochemical technique as well as
sharing his goofy sense of humor and conscientious perspective; Yeonee, for sharing her RNA-assaydevelopment knowledge, her scientific expertise, and her kind encouragement; Justin, for engaging
in intellectual debate, entertaining banter, and frank discussion; Gavin, for sharing his sense of
the big picture and open demeanor; Hern, for sharing his great knowledge of science, sports, andfine cuisine; and Allison, Andy, Louisa, George, Rebecca, Duc, Ruby, and Matt, for sharing theirtechnical knowledge and good-natured, friendly energy.
The members of the JILA shops and staff have added to the technical success of this work as
well as to an upbeat working environment. I would like to thank the JILA Instrument Specialists,
with special thanks to David Alchenberger, for his substantial assistance in the nanofabrication
processes required in our experiments, and Blaine P. Horner, Hans Greene, and Thomas P. Foote, fortheir work building various parts for the apparatuses used in the experiments; the JILA Electronics
Shop, with thanks to Carl J. Sauer, James Fung-A-Fat, and Terry J. Brown; the JILA Computing
Staff, with thanks to James D. McKown, Michael D. Paige, and James R. Raith; the JILA Supply
Office, with special thanks to Jeffrey J. Sauter, Brian C. Lynch, Randall T. Holliness, and David
C. Errickson; the JILA Scientific Reports Office; and the JILA Administrative Staff, with additional
thanks to Agnieszka K. Lynch and Maryly L. Dole.
I would also like to thank the director and deputy directors of the NSF-IGERT Optical Scienceand Engineering Program, Dana Z. Anderson, Barbara Tennis, and Ricki W. Hadow. OSEP not
only provided a source of funding for this work, but additionally provided opportunities for my
expanded learning and training in research, including opportunities such as an internship at Nikon
Instruments in Japan and extended coursework in an array of topics.
I would like to offer my deepest thanks to my Thesis Committee: Amy E. Palmer, John
P. Cumalat, Margaret M. Murnane, Matthew A. Glaser, Meredith D. Betterton, and Thomas
T. Perkins. Thank you so much for supporting me in this work.
Many friends I made before and during graduate school have influenced and enhanced myviii
doctoral experience: Ben Zhao, Elisa K. Berry, Nguyen Hoang, Holly A. Magdanz, Peter Bell,
Laura Horovitz Strashny, Heriberto Vargas, Terence ManChun Tong, Chul-Kyun Park, Po-Wei
Chen, Nicole M. Anthony, Esteban Gomez, Sean Haq, Samuel E. Wurzel, Rajiv Bhat, Samik Dasgupta, Manuel Castellanos Beltran, Kassandra Neff Castellanos, Javier von Stecher, Leila Gomez,
Ashley R. Carter, Gregory Herman, Amanda E. Carpenter, Martin M. Boyd, Justin A. Kuczynski,
Yeonee Seol, Gavin M. King, Karen L. King, Byungmoon Cho, Laura C. Sinclair, Galen C. O’Neil,
Shihkuang Tung, Ching-yu Claire Wu, Juan M. Pino, Russell Stutz, Giacomo Lamporesi, Aaron
E. Leanhardt, D. Hern Paik, Julie L. Fiore, Haemi Li, Stephanie A. Meyer, Qudsia Quraishi, Emily
Gibson, Carl J. Sauer, Kristine Abeyta, Ignacio Castellanos Beltran, Huanqian Loh, Kunio Toshimitsu, Yoko Fukuda, Norie Fujita, Janet E. McCombs, Meghana Rangan, Lora Nugent-Glandorf,
Allison B. Churnside, Vasiliy Fomenko, James Fung-A-Fat, Terry J. Brown, Maritza Sales, Laura
K. Mortimer, Matthew D. Seaberg, Michael S. Foss-Feig, Jesse C. Marcum, Rebecca K. Montange,
Genevieve Park, and Juthika Khargharia. I would like to especially thank Julie, Amanda, Meghana,
Janet, Zhao, Juthika, and Matt for their friendship and support. You have been a great source of
genuine encouragement, invaluable support, hearty laughter, and true inspiration.
Finally, I cannot thank my family enough, including Hannah Hosotani, Akiko Hosotani, Yutaka Hosotani, Suzuko Hosotani, Seigo Hosotani, Masako Hosotani, Yoko Arima, and Ronald Uno
for their support. I would like to give special thanks to my compassionate and unfailing sister who
has rallied and been there for me throughout, my caring and inquisitive mother for her deep loveof learning, and my thoughtful and warmhearted father who has inspired me and my interest inscience.

Contents

Chapter
1 Introduction1

1.1

Destabilization Dynamics of a Helicase . . . . . . . . . . . . . . . . . . . . . . . . . .

2

1.2

Real-Time Ribozyme Cleavage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

4

1.3

Overview of Thesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

52 Experimental Technique: Precision Optical-Trapping Assay
2.1

7

Nucleic-Acid Characteristics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

7

2.1.1

DNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

7

2.1.2

RNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

9

2.2

Principles of Optical Trapping . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

12

2.3

Instrument Design . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

14

2.3.1

Trapping . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

15

2.3.2

Position detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

15

2.3.3

Position control . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

16

2.3.4

Stabilization technique . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

16

Alignment, Calibration, and Characterization . . . . . . . . . . . . . . . . . . . . . .

18

2.4.1

Coarse alignment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

18

2.4.2

Trap-laser collimation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

19

2.4.3

Detection-laser focal-position optimization . . . . . . . . . . . . . . . . . . . .

19

2.4

x

2.5

2.6

2.7

2.4.4

Position calibration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

21

2.4.5

Laser-crosstalk minimization . . . . . . . . . . . . . . . . . . . . . . . . . . .

24

2.4.6

PZT-mirror calibration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

24

2.4.7

Differential pointing stability . . . . . . . . . . . . . . . . . . . . . . . . . . .

25

2.4.8

Active stage stabilization . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

25

2.4.9

Trap ellipticity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

27

2.4.10 Trap linearity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

28

2.4.11 Stiffness calibration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

30

2.4.12 Stiffness versus height . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

31

2.4.13 Sensitivity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

31

2.4.14 Stretching long DNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

33

2.4.15 Trap fiber coupling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

34

2.4.16 Trap power . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

34

2.4.17 Free-bead measurement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

34

2.4.18 DNA-tether stability . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

35

Sample Preparation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

35

2.5.1

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

35

2.5.2

Bead preparation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

36

2.5.3

Synthesis of DNA constructs . . . . . . . . . . . . . . . . . . . . . . . . . . .

37

2.5.4

DNA tethered-particle assays . . . . . . . . . . . . . . . . . . . . . . . . . . .

39

2.5.5

RecBCD tethered-particle assay . . . . . . . . . . . . . . . . . . . . . . . . . .

39

Data Acquisition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

40

2.6.1

Measuring DNA contour length . . . . . . . . . . . . . . . . . . . . . . . . . .

40

2.6.2

Employing a force clamp . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

41

Data Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

42

2.7.1

Quantifying fluctuations and conformational dynamics . . . . . . . . . . . . .

42

2.7.2

Determining statistical significance between data sets . . . . . . . . . . . . . .

43

xi
2.7.3

Quantifying distinct spikes within the conformational dynamics data . . . . .

3 RecBCD Conformational Dynamics and Mechanism
3.1

43
45

Helicase Function and Mechanism . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

45

3.1.1

Structural features common to all helicases . . . . . . . . . . . . . . . . . . .

45

3.1.2

Unwinding mechanisms of helicases . . . . . . . . . . . . . . . . . . . . . . . .

47

3.1.3

The versatility of the RecBCD enzyme . . . . . . . . . . . . . . . . . . . . . .

56

3.2

Directional Motion of RecBCD Along DNA Reveals Large Fluctuations. . . . . . .

63

3.3

The RecBCD·DNA Complex Exhibits Conformational Dynamics . . . . . . . . . . .

65

3.3.1

Instrumental noise at the level of 1 bp . . . . . . . . . . . . . . . . . . . . . .

65

3.3.2

Tailed DNA does not exhibit large fluctuations . . . . . . . . . . . . . . . . .

67

3.3.3

RecBCD bound to blunt-ended DNA does not exhibit large fluctuations . . .

67

3.3.4

Conformational dynamics are present in the absence of ATP . . . . . . . . . .

69

3.3.5

Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

69

3.4

Onset of Dynamics Coincides with Onset of RecD Motor Activity . . . . . . . . . . .

70

3.5

Modulation by Nonhydrolyzable ATP analogs . . . . . . . . . . . . . . . . . . . . . .

72

3.6

RecBCD Pin Undergoes Back-and-Forth Translocation Along DNA . . . . . . . . . .

76

3.6.1

Conformational dynamics are dependent on the DNA duplex sequence . . . .

78

3.6.2

Sequence-dependent dynamics observed at various forces . . . . . . . . . . . .

81

3.6.3

Conformational dynamics were suppressed with an interstrand cross link . . .

85

3.7

Quantifying RecBCD Destabilization of Duplex DNA . . . . . . . . . . . . . . . . . .

85

3.8

Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

91

3.8.1

Large conformational dynamics present in the absence of ATP . . . . . . . . .

91

3.8.2

RecBCD operates by dynamic destabilization and conformational changes ofthe enzyme . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

4 glmS Ribozyme Cleavage
4.1

Active Roles of RNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

92
94
94

xii

4.2

4.3

4.4

4.1.1

Ribozymes are catalytic RNA molecules . . . . . . . . . . . . . . . . . . . . .

94

4.1.2

Riboswitches detect specific metabolites . . . . . . . . . . . . . . . . . . . . .

95

4.1.3

The glmS ribozyme-riboswitch

97. . . . . . . . . . . . . . . . . . . . . . . . . .

Materials and Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 101
4.2.1

RNA:DNA hybrid constructs . . . . . . . . . . . . . . . . . . . . . . . . . . . 102

4.2.2

Protein-conjugated beads . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 104

4.2.3

RNA tethered-bead assay with BSA surface passivation . . . . . . . . . . . . 104

4.2.4

RNA tethered-bead assay with PEG surface passivation . . . . . . . . . . . . 105

4.2.5

Single-molecule data acquisition and analysis . . . . . . . . . . . . . . . . . . 106

4.2.6

Ensemble FRET experiments . . . . . . . . . . . . . . . . . . . . . . . . . . . 107

Preliminary Results . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 108
4.3.1

Cleavage of single glmS ribozymes . . . . . . . . . . . . . . . . . . . . . . . . 108

4.3.2

Cleavage of an ensemble of glmS ribozymes . . . . . . . . . . . . . . . . . . . 110

Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111

5 Conclusions and Future Directions

112

5.1

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 112

5.2

Future Directions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113
5.2.1

Further characterization of back-and-forth dynamics . . . . . . . . . . . . . . 113

5.2.2

Quantification of duplex destabilization by the helicase . . . . . . . . . . . . . 113

5.2.3

Characterization of the RecB motor and RecBCD after Chi recognition

Bibliography. . . 114

115

Appendix
A Protocols

131

A.1 RecBCD Tethered-Bead Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 131

xiii
A.2 glmS Ribozyme Tethered-Bead Assay
B Alignment and Calibrations. . . . . . . . . . . . . . . . . . . . . . . . . . 135
138

B.1 Optical-Trapping Instrument . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 138
C Data145

xiv

TablesTable
2.1

PZT mirror calibration values . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

3.1

RecBCD·DNA conformational dynamics of RecBCD bound to 50 -(dT)10 -tailed DNA
for various duplex sequences . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

25

82

Figures

Figure2.1

Force extension curves of DNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

8

2.2

DNA chemical structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

10

2.3

Major and minor grooves in DNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

11

2.4

Major and minor grooves in DNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

11

2.5

Optical-trapping assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

12

2.6

Optical layout for stabilized optical trapping . . . . . . . . . . . . . . . . . . . . . . .

17

2.7

Axial QPD signals of a free bead and stuck bead . . . . . . . . . . . . . . . . . . . .

19

2.8

Axial QPD signals of a free bead . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

20

2.9

QPD signals of a stuck bead translated laterally . . . . . . . . . . . . . . . . . . . . .

22

2.10 QPD signals of a stuck bead translated axially. . . . . . . . . . . . . . . . . . . . .

23

2.11 PZT mirror calibrations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

26

2.12 Laser differential pointing stability in three dimensions . . . . . . . . . . . . . . . . .

26

2.13 Stability with active stage stabilization . . . . . . . . . . . . . . . . . . . . . . . . . .

27

2.14 Trap ellipticity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

28

2.15 Trap linearity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

29

2.16 Stiffness calibration of d = 400 nm beads . . . . . . . . . . . . . . . . . . . . . . . . .

31

2.17 Stiffness calibration of d = 320 nm beads . . . . . . . . . . . . . . . . . . . . . . . . .

32

2.18 Trap stiffness versus height . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

32

2.19 Lateral sensitivity measurements of a stuck bead . . . . . . . . . . . . . . . . . . . .

33

xvi
2.20 Lateral sensitivity measurements of a free bead . . . . . . . . . . . . . . . . . . . . .

34

2.21 Trap power versus servo voltage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

35

2.22 Stability of a DNA tether at 6 pN . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

36

2.23 DNA substrates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

38

2.24 High-precision RecBCD optical-trapping assay . . . . . . . . . . . . . . . . . . . . . .

42

2.25 Quantifying spikes within the conformational dynamics data . . . . . . . . . . . . . .

44

3.1

Structure of the RecD protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

46

3.2

Representative core structures found in helicases and translocases . . . . . . . . . . .

48

3.3

Passive and active mechanisms for helicase unwinding . . . . . . . . . . . . . . . . .

49

3.4

The inchworm model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

54

3.5

The rolling model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

55

3.6

RecBCD-dependent homologous recombination pathways . . . . . . . . . . . . . . . .

58

3.7

RecBCD-catalyzed DNA-end-processing reaction . . . . . . . . . . . . . . . . . . . .

60

3.8

Structure of RecBCD . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

60

3.9

Structure of RecB arm . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

61

3.10 Directional motion of RecBCD . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

64

3.11 RecBCD·DNA displays large fluctuations when paused between stepping . . . . . . .

66

3.12 Distinct fluctuations over 0.07–10 Hz . . . . . . . . . . . . . . . . . . . . . . . . . . .

66

3.13 Potential sources of fluctuations in the single-molecule RecBCD assay. . . . . . . .

67

3.14 The RecBCD·DNA complex exhibits conformational dynamics . . . . . . . . . . . . .

68

3.15 Conformational dynamics of RecBCD·DNA depend on length of 50 -(dT) tail . . . . .

71

3.16 RecBCD bound to 50 -(dT)10 -tailed DNA in the presence of nonhydrolyzable ATPanalogs. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

74

3.17 Comparison of RecBCD·DNA fluctuations in the presence and absence of ADP·BeFx

75

3.18 Potential mechanisms for back-and-forth motion: Three scenarios of RecBCD·DNA
conformational dynamics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

77

xvii
3.19 Comparison of RecBCD·DNA fluctuations: RecBCD bound to AT-rich versus GCrich 50 -(dT)10 -tailed DNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

79

3.20 Sequence-dependence comparison of RecBCD·DNA bound to 50 -(dT)10 -tailed DNA .

80

3.21 Sequence-dependent conformational dynamics of RecBCD bound to 50 -(dT)10 -tailed
DNA at varying force . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

83

3.22 Sequence-dependent conformational dynamics of RecBCD bound to 50 -(dT)10 -tailed
DNA versus force . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

84

3.23 Conformational dynamics are altered with a cross-linked DNA . . . . . . . . . . . . .

86

3.24 Comparison of RecBCD bound to 50 -(dT)10 -tailed DNA with 4 different duplex DNA
sequences . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

88

3.25 Comparison of number of spikes in RecBCD bound to cross-linked and non-crosslinked DNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

90

4.1

Regulation of gene expression by a riboswitch . . . . . . . . . . . . . . . . . . . . . .

96

4.2

Reaction catalyzed by glucosamine-6-phosphate synthase . . . . . . . . . . . . . . . .

97

4.3

Structure of the glmS ribozyme . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

99

4.4

Schematic of the single-ribozyme cleavage assay . . . . . . . . . . . . . . . . . . . . . 101

4.5

Sequence of the glmS -ribozyme-containing RNA used in the RNA:DNA construct . . 103

4.6

Signal of a cleavage event as measure with a tethered-particle assay . . . . . . . . . . 107

4.7

Single-molecule measurements of glmS ribozyme cleavage

4.8

Real-time ensemble measurement of glmS ribozyme cleavage . . . . . . . . . . . . . . 110. . . . . . . . . . . . . . . 109

C.1 DNA tethers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 146
C.2 50 -(dT)10 -tailed DNA tethers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 147
C.3 RecBCD bound to blunt-ended DNA . . . . . . . . . . . . . . . . . . . . . . . . . . . 148
C.4 RecBCD bound to partially unwound DNA in the absence of ATP . . . . . . . . . . 149
C.5 RecBCD bound to 50 -(dT)6 -tailed DNA . . . . . . . . . . . . . . . . . . . . . . . . . 150
C.6 RecBCD bound to 50 -(dT)8 -tailed DNA . . . . . . . . . . . . . . . . . . . . . . . . . 151

xviii
C.7 RecBCD bound to 50 -(dT)10 -tailed DNA . . . . . . . . . . . . . . . . . . . . . . . . . 152
C.8 RecBCD bound to 50 -(dT)20 -tailed DNA . . . . . . . . . . . . . . . . . . . . . . . . . 153
C.9 RecBCD bound to 50 -(dT)10 -tailed DNA in the presence of ADP·BeFx . . . . . . . . 154
C.10 RecBCD bound to 50 -(dT)10 -tailed DNA in the presence of AMPPNP . . . . . . . . 155
C.11 RecBCD bound to 50 -(dT)10 -tailed DNA in the presence of ADP + Pi . . . . . . . . 156
C.12 RecBCD bound to 50 -(dT)10 -tailed DNA in the presence of ADP . . . . . . . . . . . 157
C.13 RecBCD bound to 50 -(dT)10 -tailed DNA in the presence of ATPγS . . . . . . . . . . 158
C.14 RecBCD bound to AT-rich 50 -(dT)10 -tailed DNA at 2 pN . . . . . . . . . . . . . . . 159
C.15 RecBCD bound to AT-rich 50 -(dT)10 -tailed DNA at 4 pN . . . . . . . . . . . . . . . 160
C.16 RecBCD bound to AT-rich 50 -(dT)10 -tailed DNA at 6 pN . . . . . . . . . . . . . . . 161
C.17 RecBCD bound to AT-rich 50 -(dT)10 -tailed DNA at 10 pN . . . . . . . . . . . . . . . 162
C.18 RecBCD bound to mixed-sequence 50 -(dT)10 -tailed DNA at 2 pN . . . . . . . . . . . 163
C.19 RecBCD bound to mixed-sequence 50 -(dT)10 -tailed DNA at 4 pN . . . . . . . . . . . 164
C.20 RecBCD bound to mixed-sequence 50 -(dT)10 -tailed DNA at 10 pN . . . . . . . . . . 165
C.21 RecBCD bound to GC-rich 50 -(dT)10 -tailed DNA at 2 pN . . . . . . . . . . . . . . . 166
C.22 RecBCD bound to GC-rich 50 -(dT)10 -tailed DNA at 4 pN . . . . . . . . . . . . . . . 167
C.23 RecBCD bound to GC-rich 50 -(dT)10 -tailed DNA at 6 pN . . . . . . . . . . . . . . . 168
C.24 RecBCD bound to GC-rich 50 -(dT)10 -tailed DNA at 10 pN . . . . . . . . . . . . . . . 169
C.25 RecBCD bound to AT cross-linked 50 -(dT)10 -tailed DNA . . . . . . . . . . . . . . . . 170
C.26 RecBCD bound to GC cross-linked 50 -(dT)10 -tailed DNA . . . . . . . . . . . . . . . . 171
C.27 RecBCD bound to Low-AT 50 -(dT)10 -tailed DNA . . . . . . . . . . . . . . . . . . . . 172
C.28 RecBCD bound to Low-GC 50 -(dT)10 -tailed DNA . . . . . . . . . . . . . . . . . . . . 173

Chapter 1

Introduction

Life is governed by enzymes, remarkable molecular devices that are central to every biochemical process. Enzymes are absolutely essential to human existence, whether they are functioning inhuman cells, in the microbiota of microorganisms living in our bodies, or in the ecosystems in which
we live. These highly-specialized molecules coordinate the patterns of development, regulation, andrepair within cells by accelerating a vast diversity of chemical reactions by 6–17 orders of magnitude[1, 2]. When enzyme function is altered, the effects can be disastrous. Malfunction or deficiencies
of a single critical enzyme can lead to disease [3].
Enzyme dynamics have been shown to be a significant factor in enzyme function [4]. To study
the conformational changes of enzymes, either in isolation or complexed with their substrates, computational and experimental studies are needed. Techniques such as X-ray crystallography, NMR
studies, and biochemical and mutational analysis have provided critical insights into the characterization of the dynamics of enzymes. The emergence of single-molecule techniques is complementary
to the aforementioned techniques in the pursuit to probe and understand the dynamics of various
enzymatic systems.
The advent of single-molecule manipulation [5, 6, 7, 8, 9] and single-molecule detection[10, 11, 12] methods have made highly-sensitive, real-time measurements of individual biomoleculesand cellular processes possible. In comparison to conventional ensemble methods, single-molecule
experiments offer several advantages [13, 14, 15, 16]. Molecular properties of inhomogeneous systems can be measured, revealing information that is often hidden by ensemble-averaged results.

2Single-molecule studies permit real-time observation of rarely-populated transients that are difficult
or impossible to measure with conventional methods, such as the study of the movement, force
generation, response to external force, and unfolding of biomolecules. In conjunction with ensemble
techniques, single-molecule techniques have been instrumental in unveiling the underlying molecularmechanisms of various biomolecular systems.
For example, single-molecule methods have permitted observation of the dynamics of numerous enzymes and provided insight into their mechanisms, as highlighted in the following cases. In one
case, the first measurements of the individual steps of a virus packaging motor in the bacteriophageΦ29 were observed with high-resolution optical tweezers [17]. Packaging of double-stranded DNA(dsDNA) by this ring ATPase was found to occur in 10 base-pair (bp) increments composed of four
2.5-bp steps via a mechanism requiring contacts primarily with the phosphate backbone [17, 18]. Ina second case, Cornish et al. used single-molecule fluorescence resonance energy transfer (FRET)
to observe the first real-time intersubunit rotations of the ribosome [19], a universal RNA-based
molecular machine that performs protein synthesis. The 30S and 50S subunits, which comprise theribosome, were found to spontaneously rotate between two conformational states shown to be theintermediate steps involved in translation. These two studies exemplify the scope of single-molecule
techniques to investigate enzyme dynamics unseen by ensemble methods.

1.1

Destabilization Dynamics of a Helicase
One area being explored by single-molecule work is the study of helicases and translocases[20]. DNA helicases are protein enzymes that translocate directionally through dsDNA substrates tocatalyze the separation of the complementary DNA strands. A DNA helicase harnesses the energy
from nucleoside 50 triphosphate (NTP) binding and hydrolysis to destabilize the hydrogen bonds
between complementary base pairs within a dsDNA molecule while undergoing translocation along
the dsDNA. The mechanism of this mechanochemical coupling is still not fully understood [21, 22].
The mechanism of one particular DNA helicase, the RecBCD enzyme, has several unresolved
questions [23]. One of the outstanding questions is its translocation step size. The step size of3
RecBCD has been debated due to seemingly conflicting studies. Rapid reaction techniques tomonitor the unwinding kinetics of a series of DNA duplexes of different lengths [24] with global fittingof the unwinding time courses yield the RecBCD “kinetic step size,” the average distance unwound
between two successive rate-limiting steps in the unwinding cycle [25], as 3.9 ± 0.5 bp, independentof temperature and ATP concentration [26]. However, crystallographic studies of similar helicases
bound to DNA in ATP-analog-bound and ATP-analog-free states suggest a 1-bp step size [27, 28, 29].
To investigate this discrepancy, we aimed to measure the physical step size of RecBCD directly
using a high-precision optical-trapping assay. In addition to the ability to measure the dynamics
of individual molecules, optical-trapping assays can explore the force dependence of unwinding andallow measurements of the energy landscape. Previous measurements of RecBCD with an opticaltrap were limited to 6-bp resolution [30]. To improve resolution, active stabilization of the sample
and laser intensity was incorporated to achieve sensitivity to 1-bp movements [31]. With thisimproved single-molecule assay, RecBCD translocation along DNA was measured. In doing so, we
observed large, multi-base-pair conformational dynamics of the RecBCD·DNA complex. The natureof these dynamics is discussed here.
We interpret the large RecBCD·DNA conformational dynamics as conformational changesof the enzyme and rapid back-and-forth translocation of the enzyme pin along the DNA, modulated by the enzyme’s nucleotide state. Five key observations support this interpretation. First,
the large conformational dynamics were present in the absence of ATP. Second, the onset of these
dynamics was coincident with previously-determined kinetic studies of the onset of unwinding activity, which demonstrate requirement of a sufficiently-long 50 DNA strand (≥ 10 nucleotides) toengage the RecD helicase [32]. Third, the conformational dynamics were reduced in the presence of
three particular nonhydrolyzable ATP analogs, ADP·BeFx , AMPPNP, and ADP + Pi . Fourth, theconformational dynamics were modulated by the GC-content of the DNA. Finally, these dynamics
were suppressed in the presence of an engineered, interstrand cross link in the DNA. Taken together,
these results suggest a mechanism in which large conformational dynamics of the RecBCD·DNA
complex, including dynamic destabilization of the DNA duplex by the helicase, are modulated by

4
the nucleotide-bound state of the enzyme.

1.2Real-Time Ribozyme Cleavage
Ribozymes are RNA molecules that have intrinsic enzyme-like activity. Their discovery [33, 34]

drastically expanded the view of RNA molecules from simply a messenger that passed genetic
information from DNA to proteins, to catalytic molecules able to function in the complete absenceof protein cofactors [33, 34]. It has since been shown that RNA performs major catalytic functionsin the ribosome [35, 36, 37] and spliceosome [38, 39], two essential ribonucleoprotein complexes incells. In addition, RNA can play a remarkable role in regulating gene expression [40]. Hundreds ofsmall noncoding RNAs, microRNAs, and endogenous small-interfering RNAs have been discoveredthat perform transcriptional and posttranscriptional regulation [41, 42, 43, 44]. As a result, RNAapplications have been developed in biotechnology and medicine [45, 46, 47], including developmentof RNA-based biosensors [48, 49, 50].
RNA-based biosensors have been proposed to offer a robust alternative to protein-based binding molecules, e.g., antibodies. In comparison to monoclonal antibodies, nucleic-acid-based biosensors carry distinct advantages, such as the ability to sense toxic target molecules, high stability, andease of synthesis and modification [49, 51, 52, 53]. Single-stranded DNA and RNA molecules havebeen shown to be able to bind target molecules with high affinity and high specificity, rivaling thebinding properties of antibodies [51, 52].
One particular naturally-occurring, RNA-based biosensor that has catalytic and gene regulatory functions is the glmS ribozyme [54]. This ribozyme functions by responding to binding of
its target ligand by undergoing self-cleavage, which then destabilizes the mRNA in cells to reducegene expression [54, 55]. The investigation of the glmS ribozyme provides insight into a mechanism for gene regulation by a ribozyme, as well as potentially aiding the development of synthetic,
RNA-based biosensors. Here, we developed a single-molecule cleavage assay of the glmS ribozyme
to measure cleavage of single ribozymes, providing demonstration of an assay that may be applied
to other nucleic-acid-based biosensors.

5

1.3

Overview of Thesis
In Chapter 2, we describe the experimental techniques in detail. First, we discuss the char-acteristics of DNA and RNA, the nucleic acids that are central elements in this thesis, nucleic-acidprocessing, catalytic activity, and precise manipulations. Next, we explain the underlying physicalprinciples of optical trapping, the instrument design involved in 1-bp stabilization, and the key
procedures for optimization of and measurements using the trap. Details of sample preparation,
data acquisition, and data analysis are described.
In Chapter 3, we explore the motion of individual RecBCD helicases with a 1.5-bp resolution
optical-trapping assay. First, we discuss background information on helicase, including structuralfeatures common to helicases, proposed models of unwinding and directed translocation, as wellas the importance of the role of RecBCD in DNA replication and maintenance in Escherichia coli.
Next, we present measurements of the conformational dynamics of RecBCD·DNA complexes. We
measure large, multi-base-pair-sized conformational dynamics that are consistent with RecBCD
back-and-forth translocation along DNA. These conformational dynamics are suppressed when aninterstrand cross link is incorporated into the DNA substrate. Furthermore, the amplitude of theconformational dynamics is shown to depend on the GC content of the dsDNA near the ssDNAdsDNA junction. The amplitude and frequency of the translocation are shown to be modulated by
the nucleotide state of the enzyme, as well as by the force applied by the optical trap.
In Chapter 4, we present single-molecule measurements of glmS ribozyme cleavage. First, we
discuss background information on RNA as a catalytic molecule and gene-regulatory element, as
well as background information on the glmS ribozyme, including its function in the cell and threedimensional structure. Next, we detail the experimental techniques of the single-molecule RNAbased biosensor. Synthesis of the detection element is discussed. Cleavage signals were measured
using video microscopy and utilization of tethering a nanoparticle to a surface via a ribozymecontaining RNA:DNA construct. Finally, we interpret the cleavage data of single glmS ribozymes
and conclude with future directions for this type of study.

6
In Chapter 5, we discuss several future prospects for study of the RecBCD enzyme.
The major results of this work are threefold: 1) Direct measurement of destabilization dynamics of the RecBCD·DNA complex that are qualified by new measurements of RecBCD boundto a cross-linked DNA substrate; 2) Modulation of the destabilization dynamics in the presenceof an ATP analog signifying altered conformational dynamics of the complex dependent on the
nucleotide-bound state of the enzyme; and 3) Development of a new single-molecule biosensor assay utilizing the glmS ribozyme. Although destabilization of DNA substrates has been inferred instudies of other helicases, the work presented here is, to our knowledge, the first real-time measurement of multiple-base-pair destabilization dynamics of a helicase·DNA complex. This measurement
was possible due to the high precision of our RecBCD optical-trapping assay, which was capableof measuring conformational changes at the level of 0.50 ± 0.05 nm (1.5 ± 0.2 bp, mean ± σ x̄ , at
∆f = 0.01–1 Hz on 993-nm DNA under a 6-pN load). The measurements reported here quantify
conformational dynamics that are greater than 4 bp on a 100-ms time scale and indicate translocation of the helicase pin relative to the DNA substrate, contrasting crystallographic studies thatsuggest discrete 1-bp translocation of superfamily-1 and superfamily-2 helicases along their DNA
substrates.
Additionally, this thesis presents a new biosensor sensitive to catalytic events of individualribozymes. The exploration of biosensors that use nucleic acids as the detection element is proving to
be a viable alternative to traditional biosensors, which utilize antibodies as the sensing domain. The
assay presented here enables self-cleavage to be measured in real time by visualizing each ribozyme
attached to a surface with a nanoparticle. This assay is versatile enough to be applicable to a varietyof nucleic-acid-based biosensors in addition to the glmS ribozyme, making it an important addition
to the single-molecule toolbox available for studying catalytic nucleic acids.

Chapter 2

Experimental Technique: Precision Optical-Trapping Assay

This chapter discusses the techniques of optical-trapping instrumentation to conduct singlemolecule studies of a helicase. To calibrate our optical-trapping apparatus, it is important tounderstand the physical properties of DNA under study (Section 2.1.1) and the underlying physical principles of optical trapping (Section 2.2). Section 2.3 discusses precision optical-trapping
instrumentation, Section 2.4 describes alignment and calibrations, and Section 2.5 discusses sample
preparation. Section 2.6 discusses the acquisition of RecBCD data and controls and Section 2.7
discusses methods of analysis.

2.1

Nucleic-Acid Characteristics

2.1.1

DNA

The maintenance and repair of deoxyribonucleic acid (DNA) is essential for eukarya, archaea,
and bacteria. DNA stores genetic information important for the development and functioning of
all known living organisms. Damage to DNA can be caused both endogenously and exogenously,
causing DNA lesions. Unrepaired lesions can lead to mutagenesis and carcinogenesis [56, 57] andcontribute to neoplastic transformation of cells, i.e., malignant tumors [58, 59, 60].
Studies of the elastic properties of DNA [61, 62, 63] serve as a basis for studies of DNA
remodeling by helicases, nucleases, nucleosome packaging enzymes, and other remodeling and repair
enzymes. B-form DNA, composed of two complementary single strands of DNA, is the most common
form of DNA found in biology. It assumes a right-handed double-helical structure of 10 base pairs

8

Figure 2.1: Force extension curves (FECs) of dsDNA and ssDNA. Reproduced from Woodside andValentine (2009) [68] with kind permission of Springer Science and Business Media. The dsDNA
FEC (black ) is well fit by a worm-like chain model, Equation 2.1 (green), whereas the ssDNA FEC(blue) is fit by a freely jointed chain model (red ). [Adapted from Wang et al. (1997) for dsDNA andSmith et al. (1996) (ssDNA).](bp) per turn [64], a pitch of 3.4 nm, and radius of 1.0 nm [65]. The bending stiffness of DNA isquantified by the persistence length [66], which depends on the molecule’s local elastic behavior. The
persistence length for B-form double-stranded (ds) DNA is ∼50 nm, whereas that of single-stranded(ss) DNA ranges between 0.8–1.4 nm at 0.1 M salt concentrations [67].
A single dsDNA molecule subjected to force shows a characteristic relationship between force
and extension, described by the force-extension curve [68] (Figure 2.1). This behavior is best fit by
the worm-like chain model, which treats the dsDNA as an elastic rod [63, 69]. A useful interpolation
formula for worm-like chains has been developed for fitting a force-extension curve [70]:
kB T
F =
Lp

" 

1
x
F
1− +
4
L K

−2

#

1 x
F
− + −
,
4 L K

(2.1)

where Lp is the persistence length, L is the contour length, and K is the elastic modulus of thedsDNA. At low forces, entropic elasticity of the polymer chain dominates, whereas at high forces,
enthalpic elasticity of the bonds in the backbone dominates.
A nucleic acid is a biopolymer composed of nucleotides. A nucleotide is composed of apentose sugar ring, a phosphate group, and either a purine or pyrimidine base that is linked to the
10 carbon on the pentose sugar (Figure 2.2). The alternating sugar and phosphate groups make up
the backbone of the nucleic acid. The phosphate group links the 30 carbon and 50 carbon of adjacent

9sugar molecules, giving single-stranded (ss) nucleic acids a polarity. The bases found in DNA andRNA are different: adenine (A), cytosine (C), and guanine (G) are found in both DNA and RNA;
thymine (T) occurs only in DNA; and uracil (U) only occurs in RNA. Base pairing occurs throughhydrogen bonding between bases. In dsDNA, adenine and thymine form an “AT” bp, which has
two hydrogen bonds, and guanine and cytosine form a “GC” bp, which has three hydrogen bonds.
The free energy to open a single base pair was determined from DNA unzipping experiments fit toa theoretical model [71], yielding best-fit values of 1.2 k B T for an AT bp and 3.4 k B T for a GC bp
at 25 ◦ C in 7 mM MgCl2 and 50 mM NaCl [72].
Proteins that bind DNA and recognize specific DNA sequences do so by interacting withpotential hydrogen-bond donor and acceptor atoms in the two grooves within the double helix, designated as the major and minor grooves [1] (Figure 2.3). The minor groove contains the pyrimidine
O-2 and the purine N-3 of the base pair, and the major groove is on the opposite side of the pair.
These grooves arise because the glycosidic bonds of a base pair are not diametrically opposite eachother [1] (Figure 2.4). The larger size of the major groove in B-DNA makes it more accessible forinteractions with proteins that recognize specific DNA sequences [1].

2.1.2

RNA

RNA is found in a variety of forms including messenger RNA (mRNA), transfer RNA (tRNA),RNA bound to proteins to form ribonucleoproteins, and even RNA molecules that adopt complex
three-dimensional structures to perform chemical reactions or regulate gene expression. The best
known type of RNA, mRNA, is single stranded and contains sequence information for protein
biosynthesis, i.e., translation. Some RNA is double stranded (e.g., some viral RNA, micro RNA, andsmall interfering RNA) and contributes to the immune response to viruses, as well as the regulationof gene expression. Transfer RNA molecules contain an anticodon region that base pairs to thecorresponding codon on mRNA during translation. Ribonucleoproteins are nucleoproteins that
contain RNA. RNA molecules that assume tertiary structures, such as ribozymes and riboswitches,
are discussed in Section 4.1.

10

Figure 2.2: Chemical structure of DNA.

11

Figure 2.3: Major and minor grooves in B-form DNA. Figure from Berg et al. (2002) [1]. The major
groove (orange) and the minor groove (yellow ) are depicted. The carbon atoms of the backbone
are shown (white).

Figure 2.4: Major and minor grooves in B-form DNA. Figure from Berg et al. (2002) [1]. Because
the two glycosidic bonds are not diametrically opposite each other, each base pair has a larger sidethat defines the major groove and a smaller side that defines the minor groove. The grooves are
lined by potential hydrogen-bond donors (blue) and acceptors (red ).

12Figure 2.5: Surface-coupled optical-trapping assay. Reprinted from Carter et al. (2009) [31] withpermission from Elsevier. A DNA molecule is held between two points, a cover slip and a trapped
bead (purple). One laser (red ) monitors the position of a fiducial mark in three dimensions (e.g.,
xfid ) to actively suppress mechanical perturbations. A second laser (green), collinear to the trap,
measures bead position (xbd ). The xstage is the distance between the DNA’s anchor point to thecover slip and the center of the optical trap.

The molecular composition of RNA is very similar to that of DNA. As mentioned in 2.1.1,
the base, uracil (U), only occurs in RNA. RNA nucleotides contain a ribose while DNA nucleotides
contain a deoxyribose. The additional 20 -hydroxyl group increases the susceptibility to spontaneoushydrolysis of the phosphodiester bonds in RNA. Some ribozyme catalytic activity involves nucleophilic attack of a 20 -hydroxyl on the adjacent 30 -phosphate group [73, 74], such as that of small
RNA ribozymes of ∼40–160 nucleotides that catalyze site-specific self-cleavage.

2.2

Principles of Optical Trapping
Optical trapping of dielectric particles, also referred to as optical tweezers, is one of the mostsensitive tools for manipulating single molecules and measuring their properties [68, 75]. An opticaltrap consists of a strongly focused light beam that can apply forces in the piconewton range tomicron-sized, neutral particles [76, 77]. In a typical surface-coupled optical-trapping assay, a DNA
molecule is attached to a surface at one end while attached to an optically-trapped bead at theother [78] (Figure 2.5).
A dielectric particle near the focus of a laser will experience a force due to the transfer ofmomentum from the radiation pressure of incident photons. Traditionally, this optical force isdecomposed into two components: a scattering force and a gradient force [70]. The scattering

13
component of the force, in the direction of light propagation, arises from momentum transfer fromthe incident photons to the particle. The gradient component of the force, in the direction of
the spatial light gradient, arises from the fact that a dipole in an inhomogeneous electric field
experiences a force in the direction of the field gradient [79]. It follows that due to balance between
the scattering force and the gradient force, the trapped particle axial equilibrium position is locatedslightly down-beam from the trap focus. The gradient trapping force arises from fluctuating dipolesin the dielectric particle induced by the laser, which interact with the inhomogeneous electric fieldat the focus [70].
In developing a theoretical treatment of optical trapping, it is useful to examine the twolimiting cases for which the force on a sphere can be calculated, the Mie and Raleigh regimes. The
conditions for Mie scattering are satisfied when the trapped sphere radius (r0 ) is much larger than
the trapping laser wavelength (λ), i.e., r0 >> λ. The forces in this regime can be calculated fromray optics [80]. For a single ray of power P hitting a dielectric sphere at an angle of incidence θ1with incident momentum per second nm P/c, the forces acting through the origin are
nm P
Fscatt =c(

T 2 [cos (2θ1 − 2θ2 ) + R cos 2θ1 ]1 + R cos 2θ1 −1 + R2 + 2R cos 2θ2

)

(2.2)

and
nm P
Fgrad =c(

T 2 [sin (2θ1 − 2θ2 ) + R sin 2θ1 ]R sin 2θ1 −1 + R2 + 2R cos 2θ2

)

,

(2.3)

where nm is the index of refraction of the medium, θ1 and θ2 are the angles of incidence andrefraction, the quantities R and T are the Fresnel reflection and transmission coefficients of thesurface at θ1 , nm is the index of refraction of the medium, and c is the speed of light in vacuum.
The conditions for Raleigh scattering are met when the trapped sphere is much smaller than
the wavelength of the trapping laser, i.e., r0 << λ. In this approximation, the optical forces can becalculated by treating the particle as a point dipole. For a sphere of radius r0 , the scattering forceis [70]Fscatt =I0 σnm
,
c

(2.4)

14
σ=

128π 5 r06
3λ4

m2 − 1m2 + 2

!2

,

(2.5)

where I0 is the intensity of the incident light, σ is the scattering cross section of the sphere, m is
the ratio of the index of refraction of the particle to the index of the medium (np /nm ), and λ is
the wavelength of the trapping laser. The scattering force is proportional to the intensity. The
time-averaged gradient force is given by
Fgrad =

2πα
∇I0 ,
cn2m

(2.6)

!

α = n2m r03

m2 − 1,m2 + 2(2.7)

where I0 is the intensity of the incident light and α is the polarizability of the sphere. The gradient
force is proportional to the intensity gradient and points up the gradient when m > 1.
For small displacements, the gradient restoring force is proportional to the offset from theequilibrium position, describing a Hookean spring with a characteristic stiffness that is proportional
to the light intensity.

2.3

Instrument Design
To enable measurement of bp-scale displacements, we needed an optical-trapping assay with

1-bp stability. A typical optical-trapping instrument cannot achieve 1-bp resolution [31]. However,many biophysical systems exhibit dynamics that are on the scale of a single base pair, including
enzymes that translocate along DNA. For instance, the 1-bp step of RNA polymerase was measured[81] using a dual-beam optical-trapping assay to reduce surface-induced noise [81, 82]. To determine
whether RecBCD takes 1-bp steps along DNA, a surface-coupled optical-trapping assay with atomicscale (∼0.1 nm) stability was developed [31]. An inverted microscope (Nikon, TE-2000) was modified
to allow trapping at the specimen plane, dynamic control of the lasers and sample, improved stability,and high-resolution detection [31, 83].

15
2.3.1

Trapping

Our essential components for optical trapping included a high-powered neodymium-doped
yttrium orthovanadate crystal laser (Spectra-Physics, Millennia IR, 10 W, λ = 1064 nm), beam
expansion optics (New Focus, f = 8 mm Aspheric Lens, and Thorlabs, Plano-Convex Lens, f =100 mm), two stabilized diode lasers (Blue Sky Research, VPSL, 50 mW, λ = 785 and 850 nm),
two-axis PZT mirrors to steer the lasers (Physik Instrumente, S330.2SL), a high numerical aperture
objective (Nikon, PlanAPO-100X-IR, numerical aperture = 1.4), a sample holder, and quadrant
photodiodes for detection (PerkinElmer Optoelectronics, YAG-444-4A with custom electronics).
The trap stiffness was controlled by an acousto-optic modulator (AOM; Isomet, 1205C), which
consists of a crystal with an optical diffraction grating that is generated by an acoustic wave. The
amplitude of the acoustic wave was modulated to control the intensity of the deflected first-orderdiffracted light, thus controlling the trap intensity over a broad range. Similarly, the intensity ofeach detection laser was controlled by separate AOMs (generally set to constant values). The AOMs
additionally served as part of feedback loops used for intensity stabilization (see Section 2.3.4).

2.3.2

Position detection

Precise, high-bandwidth position detection of the trapped polystyrene bead was achieved
through back-focal-plane (BFP) detection [84, 85, 86]. BFP detection provides the three-dimensional
position of the trapped object with nanometer-scale resolution and a bandwidth up to several MHz,
only limited by the photodiode performance and amplifying electronics [84, 85]. Laser light passing
through the specimen was collected on a quadrant photodiode (QPD) placed on the optical axis
in a position that is optically conjugate to the BFP of the microscope condenser [78]. The diodequadrants are summed pairwise, normalized, and the lateral, differential signals were derived fromthe pairs for both x and y dimensions. Normalization served to reduce the effect of intensity
amplitude changes on the QPD. The axial motion (z) was deduced from the total QPD signal,
hereon referred to as the sum signal (i.e., the total light impinging upon the four quadrants of

16
the QPD). The sum signal was offset amplified with a differential amplifier with a voltage offset.
Specifically, a reference voltage V0 was subtracted from the original sum signal Vsum and multiplied
by a gain g of 1000, i.e., g(Vsum − V0 ). The QPDs featured 250 kHz bandwidth. Detection laser
power at the QPD was ∼40 µW.

2.3.3

Position control

Lateral motion of the lasers was controlled by scanning mirrors. The lateral positions of
the optical trap and fiducial mark detection laser in the specimen plane were controlled by two
feedback-stabilized, lead-zirconate-titanate (PZT) piezoelectric scanning mirrors placed in planes
optically conjugate to the back focal plane of the objective lens. Pure rotations in this plane leadto pure translations in the image plane. This allowed dynamic computer control over the positionsof the optical trap and detection laser.
Incorporation of a piezoelectric stage allowed dynamic positioning of the sample chamber
relative to the trap and facilitated calibration. A three-axis, closed-loop PZT stage (Physik Instrumente, P517.3CD) with stage controller (Physik Instrumente, E-710.P3D) was used to position thesample chamber, calibrate the instrument, and stabilize the microscope [78].

2.3.4

Stabilization technique

To stabilize the optical-trapping assay to 0.1 nm in three dimensions, several stabilization
techniques were used [31]. An inverted optical microscope was stabilized to 0.1 nm in three dimensions [78]. Two detection lasers were used to establish a local, differential measurement reference
frame to actively stabilize the surface [78] (Figure 2.6). One detection laser monitored the trapped
bead position, while the other measured fiducial position, which was fed to the stage controllerthat moved the stage. Local detection suppresses thermal expansion and differential measurement
suppresses noise common to both lasers, such as air currents and objective lens drift [83]. Finally,
trap and detection laser noise were reduced.
Decreasing multiple types of trap laser noise was critical to achieving atomic-scale resolution

17

Figure 2.6: Optical layout for stabilized optical trapping. Reprinted from Carter et al. (2009) [31]with permission from Elsevier. Two detection lasers (DL, λ = 785 nm and 850 nm) were stabilizedusing an acousto-optic modulator (AOM) based feedback loop [78]. The trap laser (TL, purple) wasstabilized using a modified version of this method that imaged the AOM onto the fiber-couplinglens (optically conjugate planes shown in light green). All three lasers were combined with dichroics
to couple them into the objective (Obj). Mirrors in conjugate planes (light blue) to the back-focal
plane of the objective enabled independent beam steering of the three beams. We measured beadand sample position in three dimensions using back-focal-plane detection. Acronyms represent thefollowing: OI, optical isolator; PBS, polarizing beamsplitter; PD, photodiode; BS, beam sampler;
PZT, piezo-electric; QPD, quadrant photodiode; and λ/2, half-wave plate.

18[31]. Pointing, intensity, mode, and polarization noise were reduced by similar intensity stabilization
methods used for the detection lasers [78]. Pointing, mode, and polarization noise of the trapping
laser was translated into intensity noise through the combination of a single-mode, polarizationmaintaining fiber and polarizing beam splitter. The resulting intensity was sampled and actively
stabilized with a feedback loop to AOM that stabilized the intensity. The steady-state trap intensity
noise was decreased to 0.01% root mean-square at moderate laser powers [31]. With this instrument,
1-bp positional precision over the frequency range 0.03-2 Hz and sensitivity to 1-bp steps along DNA
at moderate force were achieved [31].

2.4

Alignment, Calibration, and Characterization
Alignment and calibration of the precision optical-trapping instrument requires a series ofsteps that may need several iterations (summarized in Appendix B). Also, corresponding LabVIEW
programs to run the instrument, IGOR analysis procedures, and example data records are noted(Appendix B).

2.4.1

Coarse alignment

For optimal alignment, laser beams must be level, not clipping on any optics, and parallel tothe screw holes in the optical table. Lenses and detectors should be aligned to the optical axis of
the laser beam. Optics should be positioned to minimize the beam path and placed in accordance
to standard optical imaging principles. For example, telescoping lenses are positioned for imaging
the PZT mirrors to the objective back focal plane. Beams should hit the center of the PZT mirrors.
Fiber coupling of the laser into the fiber should be adjusted such that the output from the AOM
is imaged onto the angle-cleaved fiber tip. For this system, the collimated trap laser is ∼1 inch indiameter, so care must be taken so that clipping of the beam is minimized, especially because most
optics are 1 inch in diameter. A charge-coupled device (CCD) camera is used to image the specimen
plane and should be positioned to be optically conjugate to the trapping plane.

19

Figure 2.7: Axial QPD signals of a free (purple) and stuck (blue) bead as the stage was moved in
the axial direction.

2.4.2

Trap-laser collimation

If the trap laser is not optimally collimated prior to entering the objective, then the trapping
efficiency will be dependent on the bead height above the sample surface. To adjust the collimationof the trapping laser (λ = 1064 nm) such that it keeps a trapped, free (non-tethered) bead at aconstant axial position when lowering the stage position, translate the second lens after the trap fiber
output in the axial direction, and translate the output end of the trap fiber in the axial direction.
Measure the 1064 sum signal (Vsum ) of a trapped free bead versus stage position (zstage ), scanningfrom when the bead leaves the surface to when zstage is several microns away (Figure 2.8). The
position detector sum signal measured by the QPD is proportional to the total incident intensity
at the back focal plane of the condenser, providing a means of locating the surface of the trapping
chamber and measuring the focal shift. As the bead moves through the focus of the trap laser,
the phase of light scattered from the bead changes by 180◦ relative to the unscattered light due tothe Gouy phase shift. A stuck bead signal undergoes the full shift, whereas a trapped free beadmaintains a relatively constant axial position [70] (Figure 2.7).

2.4.3

Detection-laser focal-position optimization

Steps in Section 2.4.2 were repeated using the signals for the bead detection laser (λ = 785
nm) (Figure 2.8). A coarse adjustment was performed by making the image of the bead detection

20

Figure 2.8: Axial QPD signals of a free bead. (A) The sum voltage signal measured by the 785 nm
detection laser QPD as a function of stage position. (B) The sum voltage signal measured by the1064 nm trap laser QPD as a function of stage position.

21
laser focus on the CCD camera in the same image plane as that of the trapped bead. We measured
the 785 sum signal (V785,sum ) of a trapped, free bead versus stage position (zstage ), scanning fromwhen the bead left the surface to when zstage was several µm away. Then, we translated the second
lens after the fiber output in the axial direction, and translated the output end of the fiber in theaxial direction to “flatten” the bead position signal such that it stayed constant as zstage increased.
The CCD camera was positioned in a plane conjugate to the trapping plane. We determined
the z position at which a trapped bead leaves the cover slip surface by overlaying a 785-nm z scanof a free bead with that of a stuck bead (e.g., surface-to-peak ∼340 nm). We set the CCD camera
position to be at the image plane of when a stuck bead was at that position (Figure 2.7).
We positioned the fiducial mark detection laser (λ = 850 nm) focus at ∼300 nm below thetrapping plane (i.e., below the trap focal position) for optimal sensitivity to measure sample position
relative to the trap, used for active sample stabilization. The trap position can be determined using
the aligned 785 QPD sum signal off a stuck bead. We measured the sum signal off the bead withthe stage detection laser (λ = 850 nm). The 850 nm second lens and fiber tip were axially moved
to optimize the 850 nm position of the focus (e.g., to be in line with the 785 laser focus, or, in thecase of the RecBCD assay, to be 300 nm below the 785-nm laser focus).

2.4.4

Position calibration

Position calibrations were performed to determine the bead or fiducial mark position fromthe response of the QPD detector signals of the detection lasers. To calibrate position of a fiducial
mark, the fiducial mark is moved across the detector region by the stage in x (Figure 2.9), y (Figure2.9), and z (Figure 2.10). To calibrate the position of a trapped bead, the bead is moved across thedetector region by steering the trap. During fiducial or bead movement, the detector output signal
is recorded and then plotted as a function of position. The position calibration is performed eachtime a bead or fiducial mark measurement is made.

22

Figure 2.9: QPD signals of a stuck bead translated laterally. QPD x signals are shown in orange,
and QPD y signals are shown in green. (A) 785 nm laser QPD lateral signals of a bead translated
in x. (B) 785 nm laser QPD lateral signals of a bead translated in y. (C) 850 nm laser QPD lateralsignals of a bead translated in x. (D) 850 nm laser QPD lateral signals of a bead translated in y.
(E) 1064 nm laser QPD lateral signals of a bead translated in x. (F) 1064 nm laser QPD lateralsignals of a bead translated in y.

23

Figure 2.10: QPD signals of a stuck bead translated axially. The x, y, and z signals are displayedin orange, green, and blue, respectively. (A) 785 nm laser QPD sum signal. (B) 785 nm laser QPD
crosstalk signals for the trace in (A). (C) 850 nm laser QPD sum signal. (D) 850 nm laser QPD
crosstalk signals for the trace in (C). (E) 1064 nm laser QPD sum signal. (F) 1064 nm laser QPD
crosstalk signals for the trace in (E).

24
2.4.5

Laser-crosstalk minimization

We measured the Vx , Vy , and Vz voltage signals of a stuck bead when moving the stage(varying xstage , ystage , and zstage ). To minimize the crosstalk during lateral movement (i.e., minimize
Vx versus ystage and Vy versus xstage ), we optimized the rotational position of the QPD detectors.
The crosstalk values obtained over a range of ± 200 nm for xstage and ystage region were < 2%
(λ = 785 nm), < 3% (λ = 850 nm), and < 8% (λ = 1064 nm). To minimize the crosstalk during
axial movement (i.e., minimize Vx versus zstage and Vy versus zstage ), we adjusted the lateral positionof the fiber output with the 3D-translation stage upon which it was mounted. The values obtained
over a ± 200 nm range for zstage were < 1% (λ = 785 nm), < 1% (λ = 850 nm), and < 13%(λ = 1064 nm).

2.4.6

PZT-mirror calibration

PZT mirrors were used to steer the trapping and fiducial mark lasers. The mirror controller
reads two voltage inputs (Vx , Vy ), which correspond to translational motion of the laser in thespecimen plane. Each PZT mirror was calibrated by recording the QPD signals for a grid of PZT
mirror voltage inputs [87], e.g., ± 3 V with 5 iterations per side. At each requested PZT mirror
voltage coordinate, (Vx , Vy ), the stage moved correspondingly and laterally scanned and centereda fiducial mark. The centered positions, (Xcen , Ycen ), were recorded and plotted (Figure 2.11).
The distance the laser moved per volts requested from the PZT mirror were determined for eachaxis, along with the crosstalk values. The volts-to-nm calibrations were deduced from fitting thecoordinate data along each axis, as well as from the crosstalk coordinate data. Typical calibration
values were are noted in Table 2.4.6.
We measured the calibration error by requesting a grid of coordinates to the PZT stage,scanning a fiducial mark in both x and y via moving the PZT stage, and scanning the fiducial markagain in both x and y via moving the PZT mirror. The scanned signals were compared. Here, thex and y error values for ymirror = 0 and xmirror = 0 are plotted (Figure 2.11). The error values were25
Table 2.1: PZT mirror calibration values.
Mirror calibration (nm/V)
PZT850nm PZT1064nm
x-816.8-815.14y-557.9699.68xcrosstalk-9.25-17.88ycrosstalk
11.11-13.36

typically < 10 nm. Each PZT mirror can be rotated by orienting the mirror to different screw holes
in its gimbal mount. When in an optimal position, the grid was minimally tilted along the x and y
axes.

2.4.7

Differential pointing stability

To determine the pointing stability of each laser, we calculated the differential signal between
two lasers as a function of time when monitoring a common object. For example, recording thedifferential signal between the 785 nm laser and 850 nm laser (e.g., (x785 − x850 ) versus time) whenmonitoring a stuck bead quantifies the relative pointing stability between the 785 nm and 850
nm detection lasers. For the differential measurement between the two detection lasers, the average
standard deviation was (0.05 nm, 0.06 nm) in (x, y) for 1–10 Hz (averaged 10 s). For the differential
measurement between the trap and bead detection laser, the average standard deviation measuredwas (0.13 nm, 0.13 nm) in (x, y) for 1–10 Hz (averaged 10 s) (Figure 2.12). Data were smoothedwith a gaussian filter that convolved the input 100-Hz data with normalized coefficients derived
from Pascal’s triangle at the 10th level [88].

2.4.8

Active stage stabilization

We measured the performance during active stage stabilization by recording the 785 nm signal
when stabilizing with the 850 nm laser for stage-position lock. We used a DAQ rate of 10.4 kHz,averaged 104 points (to average to 100 Hz), and set the anti-alias filter to 2 kHz. First, the 785 nm
laser was centered on a stuck bead (d ∼ 400 nm) with StageDrift_1_Beam_Multi.vi and the 850

26

Figure 2.11: PZT mirror calibrations. (A) 850 nm laser PZT mirror calibration. (B) 850 nm laser
PZT mirror calibration check. x and y error values are shown in orange and green, respectively.
(C) 1064 nm laser PZT mirror calibration. (D) 1064 nm laser PZT mirror calibration check. Colors
as in (B).

Figure 2.12: Laser differential pointing stability in three dimensions. Difference in the position
signals of a common stuck bead. (A) Differential measurements between the 850 nm and 785 nm
lasers in x (fuchsia), y (green), and z (blue). (B) Differential measurements between the 1064 nm
and 785 nm lasers in x (fuchsia), y (green), and z (blue). 100 Hz data were smoothed with binomial(Gaussian filter) smoothing with 10 smoothing operations applied [88].

27

Figure 2.13: Stability with active stage stabilization. While actively stabilizing the stage with the
850 nm laser (in-loop), the position of a stuck bead was independently monitored with the 785 nm
laser (out-of-loop). (A) x (orange), y (green), and z (blue) positions of a stuck bead as monitoredwith the 785 nm laser. (B) x (fuchsia), y (light green), and z (dark blue) positions of a stuck beadas stabilized with the 850 nm laser.

was centered on the same bead with BeamAlignment.vi (PZT 850, detector beam 850) after setting
the 850 PZT mirror to (5 V, 5 V). In ZCal2BeamStageStream_Rezero.vi, we enabled stabilization(click the ReZero-enabled button) and turned on the position clamps in x, y, and z (click those
3 buttons). We then observed the 785 (out-of-loop) signals. Positional signals for a bandwidth of
0.1–10 Hz exhibited a standard deviation of (0.1 nm, 0.1 nm, 0.1 nm) in (x, y, z) (Figure 2.13 B).
For optimal stabilization, the 785 and 850 were both run at AOM servo voltages of -9 V (although
785 servo is typically -3 V).

2.4.9

Trap ellipticity

The trap ellipticity can be determined by measuring the position of a free bead in a trap andplotting the y position versus x position. The elliptical axes of the trap should align with the x
and y axes of the PZT stage. Poor trap ellipticity alignment hampers proper calibration of the trap
stiffness. Orientation of the elliptical axes of the trap can be adjusted by rotating the post-fiber
trap laser half wave plate. A linear fit of the data should extract a slope < 0.01. Trap ellipticity
slope was measured for a bead with d = 400 nm at AOM1064 voltage of 2 V, to be –0.001 (Figure2.14).

28

Figure 2.14: Trap ellipticity. (x, y) coordinates of a trapped bead (blue) measured with the 785 nm
detection laser with 1064 AOM servo at 1 V. Data were linearly fit (black ).

2.4.10Trap linearity

To measure the region in which the trap can be approximated by a harmonic potential, we
performed a hydrodynamic drag measurement in which the displacement of the bead from the trap
center is measured as a function of the viscous drag force exerted by fluid flow. This drag force wasproduced by moving the sample at a series of constant velocities with the PZT stage. The dragforce on an object moving through a fluid at velocity ẏ is Fdrag = β ẏ, where β is the drag coefficient.
The drag coefficient for a sphere near a surface can be estimated by Faxen’s law [77]:
β=

9
1−
16



r0
h



1
+
8



6πηr0
 
  ,
r0
45 r0 4
1 r0 5
−
−
h
256 h
16 h
3

(2.8)

which depends on the bead radius r0 , the distance above the surface h, and the viscosity of theliquid η. When the trapping potential is harmonic, the drag force is described by a Hookean spring,defining the relationship between bead displacement and drag force
y=−

Fdrag
β ẏ
=−
,
ktrap
ktrap

(2.9)

where y is the displacement of the bead from the trap center. Thus, the trap stiffness may beapproximated by the force-displacement relationship. We measured the linear region of the trap±100 nm range in x and y (Figure 2.15).

29

Figure 2.15: Trap linearity. The trapping potential is described by the the Force-displacement
relationship. The trapped bead was displaced in either (A) the x or (B) the y direction by ahydrodynamic-drag force. The residual values are displayed in pN.

30
2.4.11

Stiffness calibration

The stiffness (ktrap ) of the optical trap for a trapped particle at a particular trap power canbe measured by three methods [77]: the Lorentzian Power Spectrum method, the Equipartition
Theorem method, and the Hydrodynamic Drag method. When the optical-trapping potential isapproximated as a harmonic potential, U (y) = 21 ktrap y 2 , where y is the displacement of the beadfrom the trap center, the equation of motion of a trapped particle subject to thermal motion canbe solved exactly [77]. At low Reynolds number the equation of motion isβ ẏ + ktrap y = F (t),(2.10)

where β is the viscous drag (Equation 2.8) and F (t) is a time-dependent external driving force.
The dynamics of a trapped bead undergoing Brownian motion in a parabolic potential well ischaracterized by a Lorentzian power spectrum [89]:Syy (f ) =

kB T
,
3
2π β(f02 + f 2 )

f0 =

ktrap
,
2πβ

(2.11)(2.12)

where f0 is the corner frequency. The mean square displacement of the particle is2

hy i = 2π

Z +∞Syy (f )df.
0

(2.13)

By the Equipartition Theorem, the mean square displacement is [77]:hy 2 i =

kB T
.
ktrap

(2.14)

where kB is Boltzmann’s constant and T is the temperature.
From equations (2.9), (2.13), and (2.14), three different methods were used to calibrate thektrap dependence on trap servo voltage.
We used uncoated, 400 nm diameter polystyrene beads to measure the trap stiffness using thepower spectrum, equipartition, and hydrodynamic drag methods (Figure 2.16). The three stiffness

31

Figure 2.16: Stiffness calibration of d = 400 nm beads. The three methods of equipartition theorem (orange), power spectrum (green), and hydrodynamic drag (blue), were used to calibrate thestiffness-to-volt values. The data were averaged and fit to a line. The percentage difference of themeasured values to the fit values is plotted (%). (A) Stiffness calibration in the x axis. (B) Stiffness
calibration in the y axis.

values at each servo voltage agreed to ±5%. We also calibrated the trap stiffness of streptavidincoated 320 nm diameter beads using the power spectrum and equipartition methods. The power
spectrum and equipartition methods agreed to within 10% (10%) for 400 nm (320 nm) beads. To
check the calibration error, three beads were used (Figure 2.17).

2.4.12

Stiffness versus height

We measured the trap stiffness as a function of bead height above the surface. The trap
stiffness was found to decrease by < 15% at 1 µm above the surface in y and remained constant inx.

2.4.13

Sensitivity

The sensitivity of the QPD signal from a bead or fiducial mark depends on the axial position
relative to the laser focus. The sensitivity was characterized by measuring the lateral signals of afiducial mark at a range of set axial stage positions (e.g., –1000 nm< zstage < +1000 nm). The
lateral signals were fit to a 7th order polynomial (e.g., Vy (y) = A0 + A1 y + A2 y 2 + . . . + A7 y 7 ).
The sensitivity (V per nm) can be described by the inverse of the coefficient of the linear term, A−1132

Figure 2.17: Stiffness calibration of d = 320 nm beads. The equipartition theorem (orange) andpower spectrum (green) were used to calibrate the stiffness-to-volt values. The data were averaged
and fit to a line. (A) Stiffness calibration in the x axis. (B) Stiffness calibration in the y axis. (C)The % error when the kx stiffness calibration was tested on 3 beads. (D) The % error when the ky
stiffness calibration was tested on 3 beads.

Figure 2.18: Trap stiffness versus height. (A) The trap stiffness in the x axis as measured by theequipartition method (orange) and the power spectrum method (green). (B) The trap stiffness
in the y axis as measured by the equipartition method (orange) and the power spectrum method(green). Constant trap stiffness values (black ) displayed for reference.

33

Figure 2.19: Lateral sensitivity measurements of a stuck bead. The A1 coefficients in x (orange)
and y (green) of a 400-nm diameter stuck bead as a function of stage position.

(Figure 2.19).
For a trapped, free bead, the sensitivity should not vary as a function of height above thesurface. The A1 coefficient of a 400-nm diameter trapped, free bead in water increased by 5% in
both x and y at a 1000-nm height above the surface (Figure 2.20).

2.4.14Stretching long DNA

The force versus extension relationship requires measurements of both force and extension in
the direction of the applied stretch, which lies at a variable angle θ in the x-z plane and changes
during the course of an experiment. From the geometrical relationship of these parameters to theinterferometer signals, the axial position of the bead (xbead ), the DNA extension (xDNA ), and theforce exerted by the bead on the DNA (F ) can be derived [63]. The force-extension curve forstretching a relatively long DNA molecule (e.g., L >> Lp ) can be described by the worm-like chain
model (Equation 2.1). Fitting to this model, one can extract the persistence length (Lp ) and contour
length (L), which is useful for distinguishing between beads tethered to the surface via one DNA
molecule or via multiple DNA molecules.

34

Figure 2.20: Lateral sensitivity measurements of a free bead. The A1 coefficients in x (orange) andy (green) of a 400-nm diameter free bead in water as a function of stage position.

2.4.15Trap fiber coupling

Because the trap stiffness was dynamically changed by varying the requested 1064 servo
voltage (i.e., PD1064 voltage), the fiber coupling needs to be such that the coupling efficiency is
sufficient over a range of AOM voltages. This was done by optimizing the fiber coupling of thetrapping laser at an AOM voltage of 0.6 V such that the PD1064 read > 7 V. When fiber alignmentis optimized, at a servo value of 0.9 V, the PD1064 should be > 10 V. To optimize fiber coupling, thepositioning of the laser beam was adjusted via two mirrors upbeam of the fiber and the positioningof the fiber tip was adjusted via its mount [90].

2.4.16

Trap power

The power of the optical trap prior to entering the microscope should depend linearly on the1064 nm laser AOM Servo voltage. We measured the trap power versus servo voltage, fit the datausing a linear fit, and retrieved a slope of 70 mW/V (Figure 2.21).

2.4.17

Free-bead measurement

The stability of a trapped, free bead was measured to quantify the trapping stability, independent of the PZT stage. A 320-nm diameter free bead was trapped and the QPD signal was
analyzed by calculating the integrated fluctuations. The integrated fluctuations (∆f = 0.02–10 Hz)

35

Figure 2.21: Trap power versus servo voltage.

was 0.12 nm2 and 0.02 nm2 at trap stiffnesses of 0.09 pN/nm and 0.23 pN/nm, respectively.

2.4.18

DNA-tether stability

We measured the stability of a trapped bead tethered to a surface via a DNA molecule (Figure2.22). For more data, see Section 3.3. Either a force clamp or “pseudo force clamp” (see Section
2.6) was implemented. Typical settings were a force of F = 6 pN, bead diameter size of d = 320
nm, DNA length of L = 1007 nm, data acquisition rate of 10 kHz, anti-aliasing low-pass filter of 2
kHz, in Wash Buffer (see Section 2.5.5). For these settings, we demonstrated a DNA contour lengthstability of 0.50 ± 0.01 nm (∆f = 0.1–10 Hz).

2.5

Sample Preparation

2.5.1

Fiducial mark fabrication

We primarily used two methods to make fiducial marks. For both methods, we constructed
epoxy-rigidified flow chambers (internal volume 15 µl) for enhanced stability using double-stick
tape (3M) as a spacer and 5-min epoxy (Devcon) for rigidity. For taking diagnostics of the trapping
instrument, we used a 400-nm diameter bead baked onto a cover slip.
For studying RecBCD with active stabilization we used a second method, fabrication of fiducial marks onto glass cover slips. An array of low-index glass posts of 1 µm diameter with 15 µm-

36

Figure 2.22: Stability of a DNA tether at 6 pN. (A) Contour length versus time. Data were smoothed
to 10 Hz (grey) and 1 Hz (black ). (B) Integrated fluctuations of the record in A.

spacing were fabricated on glass cover slips using hydrogen silsesquioxane and e-beam lithography[78, 90]. After nanofabrication and use in single-molecule assays (e.g., the RecBCD tethered-particle
assay, Section 2.5.5), the nanofabricated cover slips were re-used after cleaning with a 10-min piranha
etch (100 mL sulfuric acid and 15 drops hydrogen peroxide at 80 ◦ C). Cover slips were recovered(>20 times) and cleaned after use by soaking the flow chamber in water for 24 h, then boiling for
1 h (to remove the cover slip), and piranha cleaning the cover slip for 10 min. Surface chemistry
after such cleaning was compatible with single-molecule biophysics experiments.

2.5.2

Bead preparation

To prepare the beads for free-bead and tethered-bead measurements, the beads were transferred to new buffer and were monodispersed with sonication. When using protein-coated beads, it
was important to reduce the amount of free protein in solution by repeat washings which involved:i) mixing beads with 0.4 % Phos-Tween buffer, ii) centrifuging the bead-buffer solution, and iii)
removing the supernatant. After the repeat washings, the beads were reconstituted to the desiredconcentration in the desired final buffer (usually Wash Buffer). Next, the beads were checked formonodispersion with an optical microscope and sonicated until adequately monodisperse.

37
2.5.3

Synthesis of DNA constructs

DNA was prepared with polymerase chain reaction (PCR) (EMD Chemicals, KOD Hot Start
DNA Polymerase) with M13mp18 plasmid template and with one biotin-labeled primer and one
unlabeled primer for DNA for the RecBCD tethered-bead assay.

This created a blunt-ended

DNA molecule (L = 2413 nm, 993 nm) with only one biochemical linkage for attachment to thestreptavidin-coated optically-trapped bead.
To make tailed DNA we cut the 993-nm-length blunt-ended DNA with BsmBI restriction
endonuclease (New England Biolabs) at a restriction site such that the purified product had alength of L = 901 nm. We purchased DNA oligos to make tailed substrates: one with a 30 poly-dT
tail (dT = 6) and a 50 end that was complementary to the BsmBI overhang, the other with a 50
poly-dT tail (dT = 6, 8, 10, or 20) and a sequence that was complementary to the first oligo up
until the 50 end. The two oligos were annealed at 70 ◦ C, slowly cooled (4 ◦ C increments held for30 s) to room temperature, and ligated using T4 DNA ligase (Invitrogen) to the cut DNA (16 ◦ C
for 16 h, 50:1 molar ratio of tailed substrate to cut DNA). The final dsDNA length of the tailed
substrates was 908 nm. We gel purified the product and concentrations were determined, as above.
To verify that the final product was a tailed DNA we further cut an aliquot of the product with a
DrdI restriction endonuclease (New England Biolabs) at a site upstream of the BsmBI site (Figure
2.23A). This produced an 829-nm-length DNA molecule containing the biotin label, and a 78-nmlength DNA molecule with the tailed substrate. If the digestion or ligation were faulty then bands
would appear at other lengths (i.e., a 164-nm length indicated the blunt-ended DNA molecule anda 72-nm length indicated the DNA molecule with the BsmBI cut end and no ligation). There were
multiple different tailed substrates that were created (see Figure 2.23B): a GC-rich substrate, anAT-rich substrate, a substrate with a mixed sequence, and cross-linked substrates.

38

Figure 2.23: DNA substrates. Adapted from Carter (2008) [90]. (A) Three main types of DNA
were used to tether streptavidin-coated beads to the surface via either a digoxigenin-antidigoxigenin
linkage or via RecBCD bound to the surface: i) Bio-dig DNA; ii) Blunt-ended DNA; and iii) Bluntended DNA modified with noncomplementary ssDNA tails on the RecBCD-binding end. DNA
constructs were labeled with biotin (Bio) and digoxigenin (Dig) tags and contained restriction sites
as marked. (B) DNA end sequences of the (dT)-tailed DNA constructs that were ligated to create
the Tailed DNA in A. The sequence of the RecBCD-binding end of the Blunt-ended DNA is also
shown.

39
2.5.4

DNA tethered-particle assays

Our DNA tethered-particle assay was adapted from those in earlier single-molecule studies[31]. Double-stranded DNA was prepared by polymerase chain reaction using one digoxigeninlabeled and one biotin-labeled primer (Integrated DNA Technologies, Coralville, IA). These labels
enabled us to attach one end of a DNA molecule (L = 1007 nm) to a streptavidin-coated bead
(320-nm diameter; Spherotech, Lake Forest, IL) and the other end to an antidigoxigenin-coatedcoverslip. Coated coverslips were made by incubating 20 µg/mL antidigoxigenin antibody (fromsheep, Roche Applied Science, Indianapolis, IN) suspended in 0.1 M sodium phosphate buffer for 1
h in flow chambers. Bead-DNA complexes were made by incubating streptavidin-coated beads (920pM) at a 9:1 molar ratio with labeled DNA at room temperature. After washing the flow chamberwith Wash Buffer, the bead-DNA mixture was flowed into the chamber and allowed to incubate for
1 h before washing again. Coverslips were reused (>20 times) by soaking the flow chamber room
temperature water overnight and then in boiling water for ∼1 h (to remove the coverslip) and thencleaning the coverslip with a 10-min Piranha etch.

2.5.5

RecBCD tethered-particle assay

Our RecBCD tethered-particle assay was adapted from those in earlier single-molecule studies[30, 91, 92]. We used the RecD biotin-tagged RecBCD derivative (RecBCD-bio), which has a biotin
moiety near the carboxy terminus of the RecD subunit. This enzyme was shown to be functional invivo [93] and exhibited nuclease, helicase, and DNA-dependent ATPase activities in vitro analogous
to those of the wild-type enzyme [91].
This protocol is summarized below and a more detailed description is included in Appendix
A.1. First, flow chambers were assembled using the piranha-cleaned, nanofabricated post cover slip
described (Section 2.5.1). Next, bead-DNA complexes were prepared. Streptavidin-coated carboxyl
beads (Spherotech, 320-nm diameter) were sonicated, then incubated for 1 h with DNA (9:1 mol
ratio, 920 pM beads) in Wash Buffer [25 mM Tris (pH 7.5), 1 mM Mg(CH3 COO)2 , 1 mM NaCl,

40
1 mM dithiothreitol, 0.4% Tween 20 (Sigma-Aldrich, St. Louis, MO), and 3 mg/ml bovine serum
albumin [BSA; concentration is that prior to filtration through a 0.22-µm filter; Millipore, Billerica,MA)]. Concurrent to bead-DNA preparation, RecBCD surfaces were prepared. Three steps were
performed before introducing the bead-DNA complexes in the flow chamber, with the flow-chamber
washed twice with 200 µl Wash Buffer and allowed to sit for 10 min after each step: (i) biotinylated
BSA (Vector Laboratories, Burlingame, CA) at 100 mg/ml in 100 mM sodium phosphate buffer (pH= 7) was incubated within the flow chamber for 20 min at room temperature to promote adsorption,(ii) 5 mg/ml streptavidin (Vector Laboratories, Burlingame, CA) was incubated for 20 min, and(iii) 15 nM RecBCD-bio was incubated for 2 h. Finally, the bead-DNA mixture was introduced
and incubated for 40 min and then washed with Wash Buffer. Conformational dynamics of the
RecBCD·DNA complex could then be observed with these RecBCD·DNA tethers.
DNA unwinding by RecBCD was initiated by introducing Wash Buffer supplemented with
2 µM ATP, 1.1 mM Escherichia coli single-stranded binding protein (SSB) (Promega, Madison,
WI), and an oxygen-scavenging system (6 mg/ml glucose (MP Biomedicals, Solon, OH), 0.2 mg/ml
glucose oxidase (Roche Applied Science, Indianapolis, IN), 30 units/ml catalase), unless otherwise
stated. Where indicated, 2 µM ATP was substituted with 2 mM of the following compounds(purchased from Sigma-Aldrich, St. Louis, MO): adenylyl imidodiphosphate (AMPPNP), adenosine
diphosphate (ADP) with and without inorganic phosphate (Pi ), adenosine 50 -O-(3-thiotriphosphate)(ATPγS), or ADP complexed with beryllium fluoride (ADP·BeFx ).

2.6

Data Acquisition

2.6.1

Measuring DNA contour length

Positional data of the trapped bead and fiducial mark were each collected on a QPD (250 kHz
bandwidth with custom electronics; ∼40 µW laser power at detector). Each QPD detected lateralmotion from the normalized intensity difference between pairs of adjacent quadrants and axial
motion from the offset-amplified sum signal of the four quadrants. The signals were filtered with

41
an 8th-order, low-pass, elliptic filter (Frequency Devices), and digitized at either 4 or 10 kHz witha 16-bit, 20-V-range data acquisition card (National Instruments, PCI-6052E). After digitization,
calibration curves were used to convert the x, y, and z voltage signals into positional units (2.4.4).
Customized software was written in LabVIEW (National Instruments, version 7.1) on a computer(Dell Dimension 9200) with a fast processor (Intel Core2-Duo, 2.66 GHz).
To measure DNA contour length (L) we first determined the axial position of the surface bymonitoring the bead detection laser sum signal of a trapped, tethered bead as a function of zstage
as the bead was brought into contact with the cover slip surface [70]. The axial trapped-beadposition calibration was performed, and then the stage was lowered 300 nm. We located the lateral
tether point position by performing a two-dimensional elasticity procedure, which also returned thepersistence length (Lp ). Bead-DNA complexes anchored by multiple DNA molecules, as determinedby a low persistence length, were not studied. Next, we centered the fiducial-mark detection laser ona nearby fiducial mark by steering this laser with a PZT mirror and measured the volts-to-position
calibration of the fiducial mark. Finally, we measured L as a function of time by stretching it along
the y axis by positioning the PZT stage to a specified force, while moving the fiducial-tracking laser
by the corresponding distance. The sample was actively stabilized using a software-based feedback
loop with a 100 Hz update rate and 0.05 proportional gain. Software simultaneously measured
trapped bead and fiducial mark positions and calculated DNA length [63].

2.6.2

Employing a force clamp

To study RecBCD motion, we applied a “pseudo force clamp” with active stage stabilization(Figure 2.24). RecBCD unwinding of DNA causes motion of the bead relative to the trap center,creating a force on the bead by the trap. To keep the force within 15% of the initial force value,
we allowed the trapped bead to stray ± 10 nm from the clamp position, which was set to 70 nm.
When the bead position differed >10 nm from the clamp position value, the stage was triggered
to reset the bead position back to the clamp value and the fiducial detection laser was moved bya corresponding distance by a PZT mirror. Raw data were collected at either 4 or 10 kHz, filtered

42

Figure 2.24: High-precision RecBCD optical-trapping assay. Figure from Carter (2008) [90]. A DNA
molecule is stretched between the RecBCD helicase and an optically-trapped bead. One detection
laser (red ) measures the bead position (xbd ), which is used to calculate DNA length. The otherdetection laser (green) measures the position of a fiducial mark (xfid ), which is used to stabilize thesurface relative to the optical-trapping laser. This RecBCD assay achieves 0.5-nm precision in DNA
length measurements (∆f = 0.1–1 Hz).

using an 8th-order, low-pass, elliptic filter, and digitized with a 16-bit, 20-V-range data card.

2.7

Data Analysis

2.7.1

Quantifying fluctuations and conformational dynamics

To quantify the conformational dynamics of the RecBCD·DNA complex, we measured theintegrated fluctuations over the frequency bandwidth 0.07–10 Hz. First, we measured ∼200-s recordsof tethered nanoparticles (e.g., DNA contour length versus time). Next, we selected 30-s regionswith minimal low-frequency noise (e.g., regions where the average contour length was constant).
We calculated the power spectrum of 30 s of the raw data for a scaled power-spectrum function (seeAppendix B.1 (22)), a Hann window, and the largest window size possible for the time span. Next,
we calculated the square root of the definite integral of the power spectral density over a specified
bandwidth (e.g., 0.07 Hz to f ) to give the integrated fluctuations as a function of frequency, f .
Finally, we used the integrated fluctuations at 10 Hz (f = 10 Hz; e.g., integrated over the frequency
bandwidth ∆f = 0.07–10 Hz) to characterize the amplitude of the fluctuations and conformational
dynamics.
We used several criteria in the selection of our data. Although our assay featured high precision
in measuring changes in DNA contour length, each individual data record contained greater levels of43absolute position uncertainty, partly due to variations in bead size (C.V. 2.2% for 320-nm-diameter
streptavidin-coated beads). Data that were unusually noisy, designated “Type 2” noise [90], was notincluded in the analysis. Furthermore, we occasionally observed tethers that appeared to lack (dT)tails in our prepared samples of RecBCD bound to DNA with 30 -(dT)6 and 50 -(dT)10 ssDNA tails.
To increase our confidence that only molecules with properly-ligated (dT) tails were included in
our analysis, we selected contour-length records that were above a certain contour length thresholdvalue at the beginning of recording. The threshold value was approximately 903 nm for DNA tethers
that were 903 nm long.

2.7.2

Determining statistical significance between data sets

The statistical significance between data sets was determined using a t-test using the twotailed distribution and heteroscedastic assumptions. Since not all sample sizes were equal, we used
a t-test that assumed that the two population variances were different. This analysis assumes anull hypothesis that the two data sets being compared are from the same population. This t-test
measures differences between data sets (but does not test equivalence) and calculates a P -value (P ),the probability of obtaining a test statistic at least as extreme as the one observed. The P -value
serves as an indicator of the confidence in which the compared data sets originated from the same
population (e.g., P < 0.05 indicates that there is a 5% chance of rejecting the null hypothesis when
it is true).

2.7.3

Quantifying distinct spikes within the conformational dynamics data

To quantify the spikes observed within the conformational dynamics data, we developed acomputer algorithm to detect spikes. The main input of the algorithm was the spike size (δspike ),defined by the user. We typically used a spike size that was the mean standard deviation of ourcontrol data, the conformational dynamics of RecBCD bound to blunt-ended DNA, which was 0.87
± 0.08 nm for a 30-s DNA contour length (L) trace smoothed to 1 Hz with a Savitzgy-Golay window.
This corresponds to a 2.6-bp spike. The criteria for a “spike” included: i) a change in DNA length

44

Figure 2.25: Quantifying spikes within the RecBCD·DNA conformational dynamics. An example
data record of the DNA contour length of an example trace of RecBCD bound to DNA (light
blue; cross-linked DNA with 30 -(dT)6 and 50 -(dT)10 tails) with spikes identified with an automatedalgorithm (black dots). The contour length data were taken at F = 6 pN and smoothed with a
Savitzky-Golay window to 1 Hz. Here, 41 spikes were identified.

that had an absolute value that was greater than or equal to the specified spike size (∆L ≥ δspike )and ii) that the spike displayed a local maximum or minimum (see Figure 2.25). These criteria
were used because the spikes were neither uniform in size nor duration. The total number of spikes
per data trace were then noted. The computer algorithm was written in Igor Pro (WaveMetrics,
Portland, OR).

Chapter 3

RecBCD Conformational Dynamics and Mechanism

3.1

Helicase Function and Mechanism
Helicases have been identified in various prokaryotes and eukaryotes, as well as in bacterio-

phages and viruses [94, 95]. Helicases and translocases function in nearly all aspects of DNA andRNA metabolism [96]. They are involved in various biological functions, including DNA replication, recombination, and repair; transcription; translation; ribosome synthesis; RNA maturationand splicing; and nuclear export.
Helicases unwind duplex DNA, RNA, and DNA-RNA hybrid molecules. To understandthe mechanisms by which helicases function at the molecular level requires information on themechanochemical coupling that occurs during nucleic-acid unwinding. To catalyze unwinding, ahelicase cycles through a series of energetic (conformational) states driven by the binding and/orhydrolysis of NTP and subsequent release of products (NDP + Pi ) [94, 97].

3.1.1

Structural features common to all helicases

Helicases and nucleic acid translocases were originally classified into five different superfamilies(SF) defined by several conserved structural motifs [98]. However, subsequent studies indicate thatthese motifs are present in enzymes that are not helicases and instead more generally identify
nucleic-acid-stimulated NTPases [21, 96]. A development of the classification incorporates a sixth
superfamily and descriptors of directionality indicated as either type A and type B, which describetranslocation along DNA in the 30 → 50 and 50 → 30 directions, respectively [96].

46

Figure 3.1: Locations of motifs in the RecD protein. Reproduced from Dillingham and Kowalczykowski (2008) [23] with permission from the American Society for Microbiology. (A) Cartoon
representation of the three-subdomain structure of RecD. Subdomain 1 (wheat) contacts the RecC
protein. Subdomains 2 (green) and 3 (red ) are equivalent to subdomains 1A and 2A of SF1 DNA
helicases. The 50 ssDNA tail (deep purple) is shown approaching these core helicase domains. (B)Cartoon representation with the seven conserved helicase motifs indicated. (C) Surface representation colored as described above for A. (D) Surface representation colored as described above forB.

47
The conserved motifs coordinate the conversion of chemical energy to mechanical energy. They
couple NTP binding and hydrolysis to enzyme conformational changes [99]. The structural motifs
are a signature for core domains that form tandem RecA-like folds found near the NTPase catalytic
site (Figure 3.1). They feature conserved residues involved in the binding and hydrolysis of ATP
equivalent to the Walker A and B boxes of many ATPases [100] and a conserved arginine residue,
termed an “arginine finger,” required for energy coupling [101]. The core domains of SF1 and SF2
helicases, made up of subdomains 1A and 2A, are made up of a single polypeptide chain containing
two RecA-like folds (Figure 3.2). The core domains of the other superfamilies are hexameric (ordouble-hexameric) rings formed from 6 (or 12) individual RecA folds. The core domains shown in
Figure 3.2A constitute the minimal translocation motor.

3.1.2

Unwinding mechanisms of helicases

DNA helicases use the energy from NTP binding and hydrolysis to unwind DNA in a directional manner. Under physiological conditions (7.8 mM ATP, 1.04 mM ADP, and 7.9 mM Pi in
Escherichia coli [3]), the free energy of ATP hydrolysis is –47.5 kJ/mol or –19.2 k B T . The interaction between the helicase and the DNA that occurs when NTP binding and hydrolysis is coupled
to directional translocation can be described by various models.
Since different DNA helicases exhibit differences in their mechanistic characteristics, suchas in their translocation rate, processivity, directionality, step size, ATP-coupling stoichiometry,and oligomeric state, several models have been proposed to describe general helicase unwinding
mechanisms [21]. In the “active” versus “passive” models, the helicase either plays a direct orindirect role in destabilizing its double-stranded DNA (dsDNA) substrate [94, 102, 103] (Figure3.3). In the active model, the helicase directly destabilizes the dsDNA at the single-stranded-DNAdouble-stranded-DNA (ssDNA-dsDNA) junction. This model requires that the helicase possess atleast two DNA binding sites [94, 104] and can either interact directly with the dsDNA at the junction
to destabilize the dsDNA, or can act indirectly by applying a torque on the duplex region throughan NTP-induced conformational change [94, 105]. In the passive model, the helicase binds ssDNA

48

Figure 3.2: Representative core structures found in helicases and translocases. Adapted from Singleton et al. (2007) [96]. Universal structural elements involved in the binding and hydrolysis of
NTP, and the coupling of this activity to conformational changes are shown in yellow. (A) The SF1and SF2 enzymes contain a monomeric core formed from the tandem repeat of a RecA-like fold.
An NTP analog (black ) is bound at the interface of the core domains. This representative structureis the core of PcrA helicase from SF1. (B) The SF3-6 enzymes contain a core that consists of six
individual RecA- or AAA+-like domains (red ) arranged in a ring. Six nucleotide-binding pockets
are present, one at each domain interface, and four are occupied with NTP analogs (black ). This
representative structure is of T7 gene 4 protein from SF4.

49

Figure 3.3: Passive and active mechanisms for helicase unwinding. Reprinted from from Soultanasand Wigley (2001) [108], with permission from Elsevier. (A) In the passive mechanism, the helicase
does not make direct contacts with the duplex. Instead, it operates by trapping ssDNA at athermally fraying ssDNA-dsDNA junction. (B) In the active mechanism the helicase interacts
directly with the dsDNA and destabilizes the duplex, thus ‘actively’ unwinding the strands.

available when thermal fluctuations partially open the DNA at the ssDNA-dsDNA junction. These
end-fraying thermal fluctuations have lifetimes of less than 1 ms [106, 107]. A passive mechanism
requires the helicase to interact with the ssDNA using at least a single site and to translocate
unidirectionally along ssDNA toward the duplex [94].
The extent to which the helicase destabilizes the junction can be described by its “interaction
potential” [109, 110], which describes how the presence of the helicase modiÞes DNA opening. If
the position of the helicase is given by n (indexing the base to which the helicase is bound) and theposition of the ssDNA-dsDNA junction is denoted by m, then the interaction between the helicase
and the junction can be characterized by an interaction potential, U (m − n), which depends only
on the separation of the helicase and the junction along the DNA. If the helicase moves toward
increasing n, we expect n ≤ m because the helicase cannot move on the dsDNA. In the case ofpassive opening, the interaction potential is described by a hard-wall potential, U = 0 for m > n
and U = ∞ for m ≤ n, in which the helicase can inhibit DNA closing but does not otherwise affectthe dsDNA. The helicase can advance only if the dsDNA opens due to a fluctuation that increases m.
During fraying events, the helicase moves forward (n increases) to prevent DNA closing. In the caseof active opening, the interaction potential is described by a softer potential with nonzero range,

50
corresponding to helicase-assisted opening of DNA, e.g., U = 0 for m > n, U = U0 − (m − n)U0 for−N ≤ (m − n) ≤ 0, and U = ∞ for (m − n) ≤ −N , for a interaction potential of potential energythat increases by energy U0 per step for N steps, allowing the helicase to take N steps beyond thepoint m = n to facilitate opening of the dsDNA. In this example, N characterizes the distance thatthe helicase interacts with the dsDNA past the junction and U0 characterizes the amount of energy
by which the helicase destabilizes the dsDNA. Choosing an interaction potential to maximize theunwinding rate, an optimized, active DNA helicase can unwind dsDNA as fast as it translocates on
ssDNA.
A related approach to characterize active versus passive mechanisms is to use the ratio of
the rate of unwinding of dsDNA (Vun ) to the rate of translocation along ssDNA (Vtrans ) and theirdependence on force and DNA sequence [111]. An active helicase displays similar unwinding andtranslocation rates, giving a ratio of Vun /Vtrans ∼ 1. A passive helicase is slowed by the presence ofthe dsDNA fork and unwinds dsDNA at a much slower rate, resulting in Vun /Vtrans << 1. For apassive helicase, the upper bound for the rate of unwinding is given by Vun ≤ αβ Vtrans [109], where
α and β denote the opening and closing rates of the DNA. The average value of αβ is approximately
1
7

for DNA [94]. Thus, the unwinding rate of a passive helicase is at most 0.14 of its ssDNA

translocation rate.
These methods have been used to analyze the unwinding of several helicases. SF1A helicases,
PcrA, Rep, and UvrD, were found to be active helicases. Crystallographic studies of PcrA resolved
destabilization of 4-5 bp near the junction [29] and footprinting and binding studies of PcrA with
mutation of residues contacting the DNA duplex produced enzymes that were deficient in helicase
activity due to reduced affinity to duplex DNA in the absence of a nonhydrolyzable ATP analog,
but had translocation activities that were the same as those of the wild-type enzyme [112], together
showing that PcrA actively engages and distorts the duplex DNA in an ATP-dependent process.
Single-turnover kinetics experiments of Rep with its 2B domain replaced by three glycines (Rep∆2B)
showed that Rep∆2B monomers exhibit a translocation rate of 530 ± 10 nt s−1 along ssDNA andan unwinding rate of 226 ± 28 bp s−1 [at 25 ◦ C in 20 mM Mops (pH 6.5), 20 % (v/v) glycerol, 5 mM

51
2-mercaptoethanol, 200 mM NaCl, 4.2 mM MgCl2 , and 3 mM ATP] [113], resulting in a Vun /Vtransratio of 0.43, which is greater than the upper-bound limit for a passive helicase.
Several studies of UvrD obtained translocation rates of UvrD along ssDNA and dsDNA. In
a magnetic tweezer single-molecule study, UvrD exhibited an average unwinding rate of 248 ±3 bp s−1 and an average rezipping rate, hVzip i, of 298 ± 3 bp s−1 , where the rezipping eventswere interpreted as enzyme-translocation-limited closing of the two DNA strands opened during
unwinding, as hVhyb i ≈ 8hVzip i, where hVhyb i is the average DNA rehybridization rate [at 25 ◦ C in
20 mM Tris·HCl, 25 mM NaCl, 3 mM MgCl2 , 0.1 mM EDTA, 1 mM dithiothreitol, and 0.5 mM
ATP] at F = 35 pN [114], resulting in a Vun /Vtrans ratio of 0.83. In stopped-flow studies, UvrD
dimers exhibited an average unwinding rate of 68 ± 9 bp s−1 [115] and UvrD monomers exhibited anaverage translocation rate of 189.0 ± 0.7 nt s−1 [116] [at 25 ◦ C in 10 mM Tris·HCl (pH 8.3), 20 mM
NaCl, 20 % (v/v) glycerol, 4 mg/ml heparin, and 0.5 mM ATP:Mg2+ ]. In another magnetic tweezer
study, UvrD exhibited an unwinding rate of 27 ± 4 bp s−1 at F = 5 pN and two translocation rates,a monomeric UvrD translocation rate of 150 nt s−1 at F = 5 pN, and a dimeric UvrD rewinding
rate of 44 ± 3 bp s−1 at F = 5 pN [at 25 ◦ C in 25 mM Tris·HCl (pH 7.4), 50 mM NaCl, 3 mM
MgCl2 , 0.2 µg/µl BSA, 1 mM dithiothreitol, and 1 mM ATP] [117], resulting in a Vun /Vtrans ratioof 0.61 for UvrD dimers at F = 5.
SF2 helicases were also found to be active, such as the hepatitis C viral NS3 helicase (NS3h)[118, 119] and the RecQ helicase [111]. Optical tweezers experiments of NS3h unwinding of an
RNA hairpin with 52% GC-content were performed and measured an average unwinding rate of
51 ± 26 bp s−1 at F = 17 pN [118]. Interestingly, unwinding of the portion of an RNA hairpin
with 100% GC-content was 22 ± 17 bp s−1 at F = 7 pN, which was >14,000-fold faster than apassive helicase, indicating that NS3h is an active helicase. Presteady-state kinetic studies of NS3h
investigated the helicase activity dependence on base-pair stability [at 22 ◦ C in 50 mM MOPS·Na(pH 7.0), 40 mM NaCl, 5 mM MgCl2 , 5 mM dithiotheitol, 0.1% Tween 20, 1 µM SSB, 100 nM NS3h,
2.5 nM unwinding substrate, and 5 mM Mg-ATP] [119]. Formulations of a single-step interaction
potential derived by Betterton and Julicher [110] were used to model the unwinding rate versus

52
bp-stability dependences, with interaction energies per base pair of ∼1.7 kcal mol−1 (2.9 kB T per
helicase) and ∼0.9 kcal mol−1 (1.5 kB T per helicase) for unwinding of dsDNA and RNA:DNA
hybrid molecules, respectively. Comparison of the modeled and observed dependences show thatthe observed dependences are not described by the passive model, providing additional evidence
that NS3h is an active helicase.
Several ring-shaped helicases were found to be passive, including the bacteriophage T4 helicase
gp41 [120] and DnaB helicase [111]. Magnetic-trapping studies of the T4 helicase were performed
and an unwinding rate at zero force was extrapolated to be 30 bp s−1 [at 25 ◦ C in 25 mM Tris·Ac(pH 7.5), 150 mM KOAc, 10 mM MgOAc2 , 1 mM dithiothreitol, and 0.5, 1, 2, 4, or 5 mM ATP] fromunwinding rate measurements at forces ranging 2–11 pN [120]. The rezipping rate versus [ATP] was
fit retrieving a maximum rate of 400 ± 10 bp s−1 . The ratio of Vun /Vtrans was 0.08, which indicatesthat the T4 helicase is a passive helicase.
The ring-shaped bacteriophage T7 gp4 helicase was measured to be partially active [72, 119].
Optical-trapping studies of the T7 helicase measure an average unwinding rate of 29 bp s−1 (σ notstated) with an assisting force of F = 5.2 pN, an average unwinding rate of 220 bp s−1 (σ notstated) with an assisting force of F = 11.2 pN, and an average ssDNA translocation rate of 322± 62 nt s−1 (mean ± σ) [at 25 ◦ C in 20 mM Tris·HCl (pH 7.5), 50 mM NaCl, 7 mM MgCl2 , 3
mM EDTA, 0.02% Tween 20, and 2 mM dTTP] [72]. The observed force-velocity dependence isconsistent with an active unwinding model [72]. Pre-steady-state kinetic studies of the T7 helicase
measured the helicase activity dependence on base-pair stability [at 18 ◦ C in 50 mM Tris·HCl (pH7.6), 40 mM NaCl, 2 mM MgCl2 , 10% glycerol, 2 µM SSB, 100 nM hexamer, 2.5 nM unwinding
substrate, and 2 mM dTTP] [119]. Formulations of a single-step interaction potential derived by
Betterton and Julicher [110] were used to model the unwinding rate versus bp-stability dependences,with an interaction energy per base pair of ∼1.2 kcal mol−1 . Comparison of the modeled andobserved dependences indicate that the observed dependences are not described by the passivemodel, providing further evidence that the T7 helicase is an active helicase.
Sub-classes within the active and passive models propose more specific helicase mechanisms,

53
including the inchworm, rolling, thermal ratchet, and Brownian motor models. The inchworm
model was proposed based on studies of Rep and helicase III [121] and has since been applied to
PcrA [29, 122] and a refined “wrench-and-inchworm” model has been applied to UvrD [27]. In the
inchworm model, a helicase possesses two non-identical nucleic acid binding sites that bind withpolarity to the DNA substrate [94, 104, 105]. The leading site interacts with the duplex to be
unwound and can bind both the duplex and ssDNA, and the tail site binds only the unwound
ssDNA (Figure 3.4 [108]). Translocation along ssDNA with DNA unwinding occurs only throughconformational changes coupled to binding and hydrolysis of one or more NTP molecules. In thismodel, the same leading site always interacts with the duplex DNA during successive unwinding
cycles.
The rolling model was proposed based on observed allosteric effects of nucleotide cofactors
on the DNA binding properties of the Rep dimer [105, 123]. The binding energetics and relative
population distrubutions of different Rep-dimer-DNA complexes were determined in the presence
and absence of nucleotide cofactors ADP and nonhydrolyzable ATP analog β-γ-imidoadenosine
50 -triphosphate [AMPP(NH)P]. Rep-dimer-DNA complexes included half-saturated bound states,consisting of one Rep protomer bound to either a ssDNA or a dsDNA and the other Rep protomer
not bound to DNA, and fully-saturated bound states, consisting of both Rep protomers boundto ssDNA, both Rep protomers bound to dsDNA, or one Rep protomer bound to ssDNA andthe other Rep protomer bound to dsDNA. Large allosteric effects were observed; binding of ADP
favored Rep dimers in which both protomers bound ssDNA, whereas binding of AMPP(NH)P
favored simultaneous binding of both ssDNA and dsDNA to the Rep dimer. Additional support
for a rolling mechanism includes the observation that Rep can bypass regions within the 30 ssDNA
that would otherwise block the helicase if continuous sliding along the DNA were required for
DNA unwinding [124], and the observation that binding of ssDNA to one subunit of the Rep
dimer stimulates release of DNA from the other subunit, a DNA-exchange reaction that is further
stimulated by ATP hydrolysis [125].
The rolling model can apply to any homo-oligomeric helicase and requires at least two identical

54

Figure 3.4: The inchworm model. Reprinted from from Soultanas and Wigley (2001) [108], withpermission from Elsevier. A monomeric helicase is shown translocating unidirectionally along a
DNA molecule, unwinding the duplex in the process.

55

Figure 3.5: The rolling model. Reprinted from Korolev et al. (1997) [127], with permission from
Elsevier. DNA unwinding and translocation by the dimeric Escherichia coli Rep DNA helicase isshown, with the subunits colored in different colors to distinguish between their alternating binding
to the duplex as the protein moves forward. The letter (P) indicates the Rep monomer. The letters(S) and (D) indicate ssDNA and dsDNA, respectively [123].

DNA binding sites, both of which must be able to bind ssDNA as well as dsDNA (Figure 3.5). The
rolling model assumes that each protomer of the helicase binds ssDNA with a defined polarity.
At least one helicase subunit is always bound to the ssDNA at the fork, while the other subunit
is bound either to the same single strand or to the adjacent duplex region ahead of the fork. In
contrast to the inchworm model, each subunit alternates binding to the ssDNA and dsDNA. The
evidence that led to the proposal of the rolling model is also consistent with the “dimeric inchworm
model,” which is currently favored for dsDNA unwinding by UvrD dimers in vitro [21, 126].
In the thermal ratchet (a.k.a. Brownian ratchet) model, the interaction potential is characterized by a hard-wall potential [109, 110]. In this scenario, the interaction potential U = 0 for
m > n and U = ∞ for m ≤ n, requiring that the helicase does not move on the dsDNA. Thismodel describes a passive helicase mechanism. In general, the interaction between a helicase anda ssDNA-dsDNA junction has nonzero range and corresponds to enzymatically-assisted opening,making the thermal ratchet model unlikely [109]. Crystal structures reveal that several helicases
can interact with ≥ 5 bp in the duplex region of the ssDNA-dsDNA junction: PcrA [29], NS3 [128],
Rep [127], [129], Dexx box DNA helicase [130], including the interaction between RecBCD with 12
bp in the duplex region past the junction [131]. Furthermore, in the case of the hard-wall potential,

56
it was shown that the ratio of the maximum average unwinding velocity to the translocation velocity
along a junction-less DNA track is limited by the free energy required to open a base pair [109].
In the Brownian motor model, a helicase translocates directionally by undergoing alternating
cycles of tight and weak binding to an inherently asymmetric DNA substrate [110, 132, 133, 134].
The SF2 NS3 helicase was found to meet the requirements of this model, in which NS3 binds tightly
in the absence of ATP and more weakly in the presence of ATP [132, 135].

3.1.3

The versatility of the RecBCD enzyme

To study helicase mechanism, we studied the RecBCD enzyme, a superfamily-1 helicase withseveral unique characteristics. It is the most rapid and most processive helicase reported in theliterature [23], unwinding DNA with velocities of ∼1000 bp s−1 [136, 137, 138, 139]. Ensemble
assays observe an average of 36,000 bp unwound per binding event [140], while single-molecule
assays record processivities as high as 48,000 bp [136]. The RecBCD holoenzyme is substantiallyfaster and more processive than RecB or RecD alone [141] or the isolated helicase subunits of
SF1 helicases PcrA, Rep, and UvrD [96, 142]. Additionally, recent experiments demonstrated thatduring unwinding, RecBCD also has the ability to disrupt various nucleoprotein complexes (e.g.,
EcoRIE111Q , RNAP, and nucleosomes) by pushing them thousands of base pairs or rapidly ejecting
them from the DNA [143]. RecBCD recognizes and binds tightly to the ends of dsDNA molecules,
with a dissociation constant in the subnanomolar to low-nanomolar range depending on the salt
concentration and on whether the DNA ends contain short ssDNA overhangs on either the 30 - or
50 -terminated strands [137, 138, 144, 145]. The macroscopic efficiency of ATP hydrolysis is between
1.3 and 3 molecules of ATP per base pair unwound, depending on reaction conditions [137, 146].
The RecBCD enzyme plays a crucial role in chromosomal DNA replication and maintenancein certain proteobacteria, including Escherichia coli cells [23, 147, 148, 149, 150, 151, 152, 153].
This SF1 helicase salvages broken dsDNA replication forks via homologous recombinational DNA
repair and degrades unwanted linear DNA in the cell through use of its helicase, nuclease, DNA
sequence recognition, and protein recruiting abilities.

57
Homologous recombination is a relatively error-free process that can repair DNA lesions using
an intact homologous donor molecule as a template (Figure 3.6). In recombination-dependent
homologous replication, DNA replication is restarted from a detached replication fork, requiring
only a single DNA end [102, 154, 149, 155]. The central step is DNA strand exchange, which
involves a joining or “synapsis” between homologous DNA molecules. This step is catalyzed by the
RecA protein, which forms a filament on ssDNA [156, 157]. This nucleoprotein filament is the active
species in the homology search and subsequent invasion of a homologous duplex DNA [158, 159].
Prior to formation of the RecA filament, broken DNA must first be processed into ssDNA.
The RecBCD enzyme initiates the repair of dsDNA breaks by converting a blunt dsDNA
end into a duplex DNA molecule possessing a 30 -terminated ssDNA tail [162, 163]. The RecBCDcatalyzed DNA-end-processing reaction is summarized as follows [23] (Figure 3.7). In step one,
RecBCD binds tightly to a blunt (or nearly blunt) DNA end of a linear DNA duplex. In step
two, RecBCD couples the hydrolysis of ATP to DNA translocation and unwinding. The ssDNA
products are cleaved asymmetrically, with the degradation of the 30 -terminated ssDNA tail being
much more vigorous than the degradation of the complementary tail. In step three, the enzyme
continues to translocate until it pauses at a crossover-hotspot-instigator (Chi) site (χ), a cis-acting
8-nt regulatory DNA sequence serving as an initiation point for recombinational dsDNA repair[164, 165]. The Chi sequence is the ssDNA sequence 50 -GCTGGTGG-30 on the 30 DNA strand[166, 167] recognized when RecBCD approaches the 30 side of the sequence as written [168]. The
recognition of Chi dramatically alters the biochemical properties of the enzyme. In step four,
RecBCD continues to translocate, but the nuclease polarity is switched; the degradation of the 30
ssDNA tail is attenuated, whereas the hydrolysis of the 50 ssDNA tail is upregulated. After Chi
recognition, RecBCD facilitates the loading of the RecA protein onto the 30 ssDNA tail. In step five,
RecBCD repeatedly deposits RecA protomers, which act as nucleation points for filament growth
primarily in the 50 to 30 direction. In step six, the RecBCD enzyme dissociates from the DNA. The
product of the enzyme is a recombinogenic nucleoprotein complex of the RecA protein bound to
the 30 ssDNA tail with Chi at its terminus. This product is able to invade homologous duplex DNA

58

Figure 3.6: RecBCD-dependent homologous recombination pathways. Reproduced from Dillingham
and Kowalczykowski (2008) [23] with permission from the American Society for Microbiology. (A)In recombination-dependent DSB repair, a broken DNA molecule containing two (previously contiguous) DNA ends is rejoined in several steps involving recombination with an intact homologous
donor DNA (“ends-in” recombination) [160]. (B) In recombination-dependent replication, a single
DNA end is formed upon the collapse of a replication fork [161]. (C) Integration of linear dsDNA
occurs by an “ends-out” mechanism [160]. Note that in all three panels, for clarity, the location of
the Chi sequence is shown only during the initial stages of each pathway.

59
to promote the recombinational repair of a dsDNA break or to restart DNA replication.
RecBCD is composed of three heterogeneous subunits. Its crystal structure was solved to
3.1 Å in the absence of a nucleotide cofactor [131] (Figure 3.8). In the crystal structure, the DNA
substrate is unwound by 4 bp, with each ssDNA tail inserted into a different entry tunnel in thecomplex. As RecBCD translocates, DNA enters as duplex, is split into two nascent ssDNA strands
that are channeled through these tunnels, and are presented to the nuclease domain at the rear.
Each of the three subunits contains important structural and biochemical features in the complex.
RecB, a DNA-dependent ATPase [169], has an “arm” that stretches to contact the incoming duplex
DNA ∼12 bp ahead of the junction [131] (Figure 3.9), a motor domain (containing core domains,
Section 3.1.1) that undergoes translocation along DNA in the 30 → 50 direction [170], and a Cterminal domain that has a 30 and 50 DNA strand endo- and exonuclease role [171, 172, 173, 174] aswell as a RecA recruitment role [175, 176]. RecC, a catalytically-dead helicase-nuclease [131, 177]
acting as a core scaffolding protein in RecBCD, forms one large hole that contacts domain 2B of
RecB, two smaller tunnels that accept the 30 - and 50 -terminated ssDNA strands, and protein-protein
contacts with RecD, serving as a structural interface between RecB and RecD. RecC contains a dualmethionine “pin” that acts as a wedge to split the duplex at the ssDNA-dsDNA junction [131] (seeFigure 3.8) and has exhibited evidence of housing a “Chi-scanning site” [178, 179]. RecD, a ssDNAdependent ATPase [141, 180], has a motor domain that undergoes translocation along DNA in the
50 → 30 direction [141]. The two motor domains of opposite directionality make RecBCD a “bipolar”
helicase [141, 181].
Previous single-molecule investigations on RecBCD directly measured DNA unwinding andin doing so, observed pausing and slipping on the DNA, events unmeasurable by ensemble methods.
RecBCD was observed to pause at Chi sites and change unwinding velocities preceding and following
Chi recognition [182, 183, 184]. In addition, RecBCD switched lead motors from RecD to RecB
before and after Chi [139]. Higher-resolution studies (∼6 bp at 1 Hz) observed force-dependent,
Chi-independent pausing and backsliding of the enzyme [30].
One of the outstanding questions regarding the RecBCD helicase is its stepping mechanism.

60

Figure 3.7: The RecBCD-catalyzed DNA-end-processing reaction. Reproduced from Dillingham
and Kowalczykowski (2008) [23] with permission from the American Society for Microbiology. (1)RecBCD (light blue) binds to the end of a blunt or nearly blunt duplex DNA. (2) RecBCD couples
ATP hydrolysis to its helicase activity. The ssDNA products are asymmetrically cleaved. (3)RecBCD translocates until it pauses at and recognizes Chi, during which the enzyme’s biochemical
properties are altered (pink ). (4) RecBCD continues to translocate, but with switched nucleasepolarity. After Chi, RecBCD facilitates RecA (yellow ) loading onto the 30 ssDNA tail. (5) RecBCD
deposits RecA protomers, which act as nucleation points for filament growth. (6) The RecBCD
enzyme dissociates from the DNA.

Figure 3.8: Cutaway view showing the channels through the RecBCD·DNA complex. Reprinted
from Singleton et al. (2004) [131], with permission from Macmillan Publishers Ltd: Nature, Copyright 2004. The cutaway surface of subunits RecB (orange), RecC (blue), and RecD (green) are
shown. RecBCD is bound to DNA (purple). Numbers refer to the domains of the appropriate
subunits. Domains 2 and 3 of RecD are equivalent to canonical helicase domains 1A and 2A.

61

Figure 3.9: The RecB arm. Reproduced from Dillingham and Kowalczykowski (2008) [23] withpermission from the American Society for Microbiology. Shown is a close-up view of the “arm”
structure (red ) formed by an auxiliary subdomain (subdomain 1B) of RecB. The arm contacts
duplex DNA ahead of the translocating enzyme. A conserved patch of residues in close proximity
to the minor groove of the DNA substrate (black semitransparent model) is shown (yellow ). The
RecC protein is in the foreground (blue). The 50 ssDNA tail is pointing toward the viewer througha hole in the RecC protein.

62
Structural and biochemical studies report different step-size values. X-ray crystallographic studiesof the SF1B Deinococcus radiodurans RecD2 helicase infer a 1-bp step size from structural analysis
of RecD2 bound to DNA with and without a nonhydrolyzable nucleotide analog bound [28]. Presteady-state single-turnover DNA-unwinding studies measure a “kinetic step size” of 4 ± 1 bp
[24, 185], where the kinetic step size is defined as the average number of base pairs unwound
between two successive rate-limiting steps in the unwinding cycle [25, 116, 186]. The kinetic step
size is the step size calculated from translocation assays that observe the final product or step of
the reaction of helicase translocation (e.g., completely unwound ssDNA) of an ensemble of helicases
in solution. The kinetic step size is not necessarily the same as the chemical step size, the numberof base pairs unwound per ATP molecule hydrolyzed; e.g., if two or more ATP molecules are boundand hydrolyzed in rapid succession, followed by a slower kinetic process, then each kinetic step will
be larger than the chemical step. The observed kinetic step size can differ from the mechanical step
size in that it can be larger if the enzyme pauses or slips, but it can also be smaller if there are
two or more kinetic processes with similar rates that occur during a mechanical step [21, 25, 26].
To address this step-size discrepancy, records of individual RecBCD enzymes unwinding DNA were
measured using a precision optical-trapping instrument [187].
The unwinding records of RecBCD displayed large, multi-base-pair conformational dynamics
of the RecBCD·DNA complex between steps that we interpret as rapid back-and-forth translocations of RecBCD’s dual-methionine pin along the DNA, modulated by the nucleotide state of the
enzyme. Here, quantification of the conformational dynamics and interpretations of the dynamics
are discussed. Several key observations were made. First, we observed large conformational dynamics in the absence of ATP, indicating that the large conformational dynamics do not originate
from enzyme interaction with a nucleotide. Second, the amplitude of the conformational dynamics
were dependent on interactions with the end of the 50 DNA strand, which interacts with the RecD
subunit, suggesting that the large conformational dynamics are modulated by enzyme interactions
with the 50 DNA strand. Together with previously measured biochemical data [32], our observations
indicate that the onset of large conformational dynamics is coincident with the onset of unwinding

63
activity by the RecD motor. Third, the conformational dynamics were altered when changing the
GC-content of the dsDNA sequence upstream of the ssDNA-dsDNA junction, indicating interaction
between the helicase and dsDNA region. Fourth, the dynamics were suppressed in the presenceof an engineered, interstrand cross link in the DNA, consistent with translocations of the enzyme
pin back and forth along the DNA. Fifth, the dynamics were reduced by the nonhydrolyzable ATPanalog, adenosine diphosphate complexed with beryllium fluoride (ADP·BeFx ), suggesting that theATP-bound state of the enzyme induces a tightly-bound state of the complex. These observations
signify the first quantification of previously-unseen, real-time measurements of active destabilization
dynamics of the RecBCD·DNA complex and lend insight into the helicase unwinding mechanism of
SF1 helicases.

3.2

Directional Motion of RecBCD Along DNA Reveals Large Fluctuations
Experiments to measure directional motion of RecBCD were carried out by tethering a dsDNA

molecule between a RecBCD molecule bound to a glass surface and polystyrene bead (Figure 2.24)
held with an actively-stabilized optical-trapping microscope [187]. Data were taken at 6 pN of force,
unless otherwise noted. With this RecBCD optical-trapping assay, we measured DNA contour length(L) as RecBCD unwound the double-stranded (ds) DNA (Figure 3.10) [90, 187]. Unwinding of thedsDNA caused a decrease in L. RecBCD motion (N = 17) displayed pauses and changes in velocity[90, 187] (Figure 3.10) as had been observed in previous experiments [30]. Records displayed discrete
forward and backward steps, which appeared as directional translocations of the enzyme separated
by pauses in movement (Figure 3.10). To quantify the steps, sections of the 1-Hz smoothed data
were fit with an automated step-finding algorithm [188] to determine the average value of L for eachplateau and its corresponding dwell time [90].
Unexpectedly, the measurements of RecBCD·DNA motion paused between steps (Figure 3.11)
exhibited large-scale fluctuations that were significantly larger than the measured noise floor of
the optical-trapping instrument (Section 3.3). The fluctuations were characterized by calculating
the power spectral density (PSD) of the DNA contour length time courses and determining the64

Figure 3.10: Directional motion of RecBCD. Adapted from Carter (2008) [90] and Carter etal. (2011). (A) DNA contour length (L) decreases as RecBCD unwinds the DNA substrate. Traces
were taken at 6 pN of force, 2 µM ATP, and smoothed with a Savitzky-Golay window to 10 Hz(light green) and 1 Hz (dark green). (B) Enlarged region (pink box ) displays forward translocation.
(C) Backward translocation was also observed. Data were fit with a step-finding algorithm (black ;line and arrows) [188].

65
integrated fluctuations (see Section 2.7), which was calculated from a frequency of 0.07 Hz. The
PSD of the records of RecBCD paused between steps (Figure 3.12) indicated that the fluctuations
occur over a frequency range of 0.07–10 Hz, which also encompasses the average forward stepping
frequency of 0.62 s−1 [187]. At 10 Hz, the mean value of the integrated fluctuations was 1.39 ± 0.02
nm (mean ± σ x̄ ) [90] for RecBCD paused between unwinding steps.

3.3

The RecBCD·DNA Complex Exhibits Conformational Dynamics
The potential sources of fluctuations in our RecBCD assay can be categorized as follows(Figure 3.13): (I) motion of the biotin-streptavidin surface attachment, (II) motion of the RecBCD
biotinylated attachment linker, (III) conformational changes of RecBCD, and (IV) translocationsof RecBCD relative to the DNA substrate. To examine the first two potential sources of noise, wemeasured a series of controls including DNA tethered to the surface in the absence of RecBCD and
DNA tethered to the surface bound to RecBCD .

3.3.1

Instrumental noise at the level of 1 bp

To demonstrate 1-bp stability of the DNA optical-trapping assay, we measured the instrumental noise using two types of DNA tethers in the absence of RecBCD. The first type was comprised of a 1007-nm dsDNA molecule with a 50 -biotin label attached to a streptavidin-coated bead
and a 50 -digoxigenin label attached to the antidigoxigenin-coated glass surface. The instrumental
noise was quantified with the same analysis methods used to quantify the fluctuations of RecBCD
paused between steps (Section 2.7). Because we observed the greatest conformational dynamics
of RecBCD·DNA as shown in the PSD in the frequency bandwidth 0.07–10 Hz (Figure 3.12), wequantified instrumental noise in this bandwidth. At 1 Hz, the first type of DNA tether exhibiteda mean integrated fluctuations value of 0.285 ± 0.011 nm (0.84 ± 0.03 bp, mean ± σ x̄ ) (Figure3.14), N = 10 (see Appendix C). At 10 Hz, the mean value of the integrated fluctuations was
0.502 ± 0.012 nm (1.49 ± 0.03 bp) (Figure 3.14), N = 10 (see Appendix C). The mean integrated
fluctuations data demonstrate the capability of our optical-trapping instrument to measure 1-bp

66

Figure 3.11: RecBCD·DNA displays large fluctuations when paused between stepping. Adaptedfrom Carter (2008) [90]. DNA contour length (L) decreases as RecBCD unwinds the DNA substrate.
Traces were taken at 6 pN of force, 2 µM ATP, and smoothed with a Savitzky-Golay window to 10
Hz (light green) and 1 Hz (dark green). Enlarged area (pink box ) is an example of RecBCD paused
between stepping.

Figure 3.12: Distinct fluctuations over 0.07–10 Hz. Representative power spectral density records of
RecBCD paused between stepping (green), RecBCD bound to the middle of DNA in the absence ofATP (red ), RecBCD bound to the end of blunt-ended DNA (blue), and DNA directly attached to asurface (black ). Grayed region indicates the frequency bandwidth over which large conformational
dynamics of the RecBCD·DNA complex occur.

67

Figure 3.13: Potential sources of fluctuations in the single-molecule RecBCD assay. Adapted from
Carter (2008) [90] and Carter et al. (2011). Fluctuations in the DNA contour length may be causedby (I) the biotin-streptavidin surface attachment, (II) the RecBCD biotinylated attachment linker,(III) conformational changes of RecBCD, and (IV) translocations of RecBCD relative to the DNA.

(0.338 nm; B-form DNA) changes in DNA contour length (∆f = 0.07–2.3 Hz).

3.3.2

Tailed DNA does not exhibit large fluctuations

In a second study of DNA tethers in the absence of RecBCD, a similar DNA construct wasused with the exception that it contained noncomplementary ssDNA tails on the digoxigenin-labeledend (“dT10 control,” Figure 2.23). The DNA end attached to the streptavidin-coated bead contained
a 50 -biotin label. The DNA end attached to the antidigoxigenin-coated glass surface had a 30 -(dT)6
ssDNA tail and a digoxigenin-labeled 50 -(dT)10 ssDNA tail. The value of the integrated fluctuations
at 10 Hz was 0.57 ± 0.03 nm (1.7 ± 0.1 bp, mean ± σ x̄ ) (Figure 3.14), N = 5 (see Appendix C),
slightly higher than that of the non-tailed DNA data, which was 0.50 nm. Only a very slightamount of noise of 0.07 nm (0.2 bp) was added with the addition of the ssDNA dT tails. This study
demonstrates that the presence of a 10-nt ssDNA region does not greatly alter the fluctuations in
our optical-trapping assay.

3.3.3

RecBCD bound to blunt-ended DNA does not exhibit large fluctuations

To examine whether the attachment of RecBCD to the surface via a biotin-linker was creating
the multi-base-pair fluctuations, we measured the fluctuations of RecBCD bound to the end of ablunt-ended DNA molecule (“Blunt end,” Figure 2.23). At 10 Hz, the mean value of the integrated

68

Figure 3.14: The RecBCD·DNA complex exhibits conformational dynamics. (A) Representative
data of the DNA contour length versus time with corresponding histograms for tethers of DNA
attached to a surface directly (black ), DNA with 30 -(dT)6 and 50 -(dT)10 ssDNA tails attached toa surface (olive), RecBCD bound to the end of a blunt-ended DNA molecule (blue), and RecBCD
bound to the middle of a partially-unwound DNA molecule in the absence of ATP (red ). Records
were taken at F = 6 pN and smoothed with a Savitzky-Golay window to 10 Hz (light) and 1 Hz
(dark ). (B) Average power spectral density. (C) Average integrated fluctuations of the PSD data.
(D) Integrated fluctuations at 10 Hz (∆f = 0.07–10 Hz, *P < 0.02, 5 ≤ N ≤ 10). All error bars
denote standard error of the mean.

69
fluctuations was 0.82 ± 0.07 nm (mean ± σ x̄ ) (Figure 3.14), N = 5 (see Appendix C) that, althoughhigher than that of the measured instrumental noise, which was 0.50 nm, was well below that of
RecBCD paused between steps, which was 1.39 nm. RecBCD bound to blunt-ended DNA was themain control used for quantification of the noise level of the RecBCD optical-trapping assay.

3.3.4

Conformational dynamics are present in the absence of ATP

To test whether the large conformational changes were present in the absence of ATP, we measured RecBCD bound to the middle of DNA in the absence of ATP. RecBCD was allowed to unwind∼1000 nm of a 2413 nm DNA in the presence of ATP and was stalled in the middle of the DNA

by flushing WB buffer (∼50x flow-cell volume) without ATP through the sample. Measurementsof the stalled complex did not exhibit directional motion, but did exhibit conformational dynamics
similar to those of RecBCD paused between steps [187]. At 10 Hz, the mean value of the integrated
fluctuations of the conformational dynamics was 1.47 ± 0.11 nm (mean ± σ x̄ ) (Figure 3.14), N = 5(see Appendix C), which was equivalent within error to that of RecBCD conformational dynamics
between steps, which was 1.39 ± 0.02 nm (mean ± σ x̄ ) [90]. Similarity of these data was expected
due to the unlikelihood of RecBCD interactions with ATP during the pauses between steps at the
low concentrations of ATP used (e.g., 2 µM).

3.3.5

Summary

These observations indicate that the fluctuations of RecBCD paused during unwinding areindeed conformational dynamics of the RecBCD·DNA complex. Records of the DNA tethers indicatethat the large fluctuations do not originate from instrumental noise. Records of RecBCD boundto blunt-ended DNA eliminate motion of the biotin-streptavidin surface attachment and motion
between RecBCD and its biotin tag as the sources of the large fluctuations. Consequently, thesecontrol experiments suggest that the large conformational dynamics of the RecBCD·DNA complex
are caused by conformational changes of RecBCD and/or translocations of the RecBCD pin relative
to the DNA.

70

3.4

Onset of Dynamics Coincides with Onset of RecD Motor Activity
We next investigated whether the large conformational dynamics of the RecBCD·DNA com-plex coincided with interaction between the RecD motor and 50 ssDNA. Equilibrium binding studies[189, 190] and structural work [191] suggest that RecBCD is able to unwind 4–6 bp of a blunt-ended
DNA molecule in the absence of ATP in a Mg2+ -dependent manner. RecBCD was shown to bind
optimally to duplex DNA with 30 -(dT)6 and 50 -(dT)m tails with m ≥ 10 nt (at 25 ◦ C in 200 mM
NaCl) [190]. Furthermore, RecBCD unwinding studies of DNA possessing 30 -(dT)6 and 50 -(dT)mtails are described by a sequential n-step mechanism with h additional kinetic steps required prior toinitiation of DNA unwinding, where h = 3.2 ± 0.1 for m = 6 and h = 0 for m = 10, indicating thatengagement of the RecD motor with the 50 ssDNA is required for RecBCD to initiate unwinding of
blunt-ended DNA [32].
To determine if conformational dynamics of individual RecBCD·DNA complexes are concurrent with the onset of unwinding activity observed in the RecBCD unwinding studies, we probed
the effect of RecD-ssDNA engagement on the conformational dynamics in the absence of ATP using
30 -(dT)6 and 50 -(dT)m DNA constructs, with m = 6, 8, 10, or 20 nt (Figure 3.15). The mean valuesof the integrated fluctuations at 10 Hz of RecBCD bound to 50 -(dT)6 -tailed DNA and 50 -(dT)8 -tailed
DNA were 0.83 ± 0.05 nm (mean ± σ x̄ ), N = 8 (see Appendix C) and 0.89 ± 0.09 nm (mean ±σ x̄ ), N = 5 (see Appendix C), respectively, both of which were equivalent within error to the meanvalue of the integrated fluctuations at 10 Hz of RecBCD bound to blunt-ended DNA (0.82 ± 0.07nm). The mean value of the integrated fluctuations at 10 Hz of RecBCD bound to 50 -(dT)10 DNA
was 1.43 ± 0.09 nm (mean ± σ x̄ ), N = 12 (see Appendix C), which was equivalent within error toboth that of RecBCD bound to 50 -(dT)20 DNA, which was 1.39 ± 0.20 nm (mean ± σ x̄ ), N = 5(see Appendix C), and to that of RecBCD paused between unwinding steps (1.47 ± 0.11 nm).
The combination of these results has two implications. First, the statistical equivalence of the50 -(dT)6 and blunt-ended data complements previous studies that indicate that RecBCD undergoes
additional kinetic steps prior to unwinding when bound to blunt-ended [24, 26, 185] and twin-(dT)6 -71Figure 3.15: Conformational dynamics of RecBCD·DNA depend on length of 50 -(dT) tail. (A)Representative data of the DNA contour length versus time with corresponding histograms fortethers of: RecBCD bound to blunt-ended DNA (blue); RecBCD bound to DNA with a 30 -(dT)6
and 50 -(dT)n ssDNA tails, with n = 6 (yellow ), 8 (purple), 10 (brown), or 20 (green) nt; andRecBCD bound to the middle of a partially-unwound DNA molecule (red ). Records were taken
at F = 6 pN and smoothed with a Savitzky-Golay window to 10 Hz (light) and 1 Hz (dark ). (B)Average power spectral density. (C) Average integrated fluctuations. (D) Integrated fluctuations at
10 Hz (∆f = 0.07–10 Hz, *P < 0.05, 5 ≤ N ≤ 12). All error bars denote standard error of the
mean.

72
tailed [32] DNA constructs. Moreover, because RecBCD has been shown to bind optimally to a30 -(dT)6 -tailed DNA end [190] that spans the RecB subunit [191], the lack of large conformational
dynamics of RecBCD bound to twin-(dT)6 -tailed DNA constructs suggests that the large conformational dynamics of the RecBCD·DNA complex do not originate from interactions between RecB
and the 30 -(dT)6 tail.
Second, the statistical equivalence of the 50 -(dT)10 , 50 -(dT)20 , and fully-threaded RecBCD
data suggest that the minimal requirement for large conformational dynamics of the RecBCD·DNA
complex is met with the 50 -(dT)10 -tailed DNA. Thus, the dependence of the magnitude of the conformational dynamics on the length m of the 50 -(dT)m tail suggests that large-scale conformational
dynamics are required to initiate binding interactions between the RecD motor and the end of the
50 DNA, which then initiate directional translocation. Since the large conformational dynamics arepresent when RecBCD is bound to 30 -(dT)6 and 50 -(dT)10 tailed DNA, we used these tailed DNA
to study RecBCD·DNA in its highly conformationally-dynamic state with a predetermined DNA
contour length to simulate RecBCD paused in the middle of DNA after unwinding, which did nothave a predetermined DNA contour length.

3.5

Modulation by Nonhydrolyzable ATP analogs
To investigate whether the conformational dynamics of RecBCD bound to 30 -(dT)6 and 50 -(dT)10 tailed DNA substrates were altered upon ATP binding, we studied this RecBCD·DNA complex in the presence of various nonhydrolyzable ATP analogs. The conformational dynamics of
RecBCD bound to 30 -(dT)6 and 50 -(dT)10 tailed DNA substrates displayed various changes in conformational dynamics as dependent on the nonhydrolyzable ATP analog present (Figure 3.16).
The conformational dynamics were significantly reduced in the presence of three nonhydolyzable ATP analogs: adenosine diphosphate complexed with beryllium fluoride (ADP·BeFx ), adenylyl
imidodiphosphate (AMPPNP), and adenosine diphosphate with inorganic phosphate (ADP + Pi ).
The most dramatic reduction was observed when RecBCD·DNA was in the presence of ADP·BeFx(Figures 3.16 and 3.17), which has been shown to act as an ATP analog in structural and kinetic

73
studies of myosin [192, 193, 194]. Conformational dynamics of the RecBCD·DNA complex in thepresence of 2 mM ADP·BeFx , 2 mM AMPPNP, and 2 mM ADP + Pi were reduced to mean valuesof the integrated fluctuations at 10 Hz of 0.91 ± 0.05 nm (mean ± σ x̄ ), N = 9 (see Appendix C), 1.07
± 0.09 nm (mean ± σ x̄ ), N = 5 (see Appendix C), and 1.03 ± 0.14 nm (mean ± σ x̄ ), N = 5 (see Appendix C), respectively. The ADP·BeFx data are equivalent within error to those of RecBCD boundto blunt-ended DNA (0.82 ± 0.07 nm) suggesting, when ADP·BeFx is bound to the RecBCD·DNA
complex, that RecD interactions with the 50 -DNA tail are restricted and that RecBCD·DNA may
only be able to access the conformational states that are available when RecBCD is bound to bluntended DNA. The conformational dynamics in the presence of ADP·BeFx , AMPPNP, and ADP + Pi
were statistically significant from the data of RecBCD·DNA in the absence of ATP analogs, with
P -values (see Section 2.7) of P < 0.0002, P < 0.02, and P < 0.05, respectively. These significant
decreases in the conformational dynamics suggest that when a nucleotide is bound to one or bothof the ATP-binding clefts of RecBCD, the enzyme adopts a conformation in which the contacts
between the enzyme and DNA are less dynamic, thus implying a tighter interaction between the enzyme and DNA. The change in the conformational dynamics of RecBCD bound to 50 -(dT)10 -tailed
DNA suggests that binding of ADP·BeFx , AMPPNP, and ADP + Pi to RecBCD modulates theselection of one or more particular conformational states of the RecBCD·DNA complex.
Conformational dynamics of RecBCD bound to 30 -(dT)6 and 50 -(dT)10 tailed DNA were notsignificantly altered in the presence of two other ATP analogs (Figure 3.16), ADP and adenosine 50 O-(3-thiotriphosphate) (ATPγS). RecBCD·DNA conformational dynamics were unchanged withinerror in the presence of 2 mM ADP and displayed a mean value of the integrated fluctuations at
10 Hz of 1.26 ± 0.10 nm (mean ± σ x̄ ), N = 5 (see Appendix C). When RecBCD was bound to30 -(dT)5 and 50 -(dT)10 tailed DNA in the presence of 2 mM ATPγS, the integrated fluctuations at
10 Hz were also unchanged within error and had a mean value of 1.29 ± 0.15 nm (mean ± σ x̄ ), N
= 7 (see Appendix C). The

74

Figure 3.16: RecBCD bound to 50 -(dT)10 -tailed DNA in the presence of nonhydrolyzable ATPanalogs. (A) Representative data of the DNA contour length versus time with corresponding histograms for tethers of RecBCD bound to blunt-ended DNA (blue) and RecBCD bound to DNA with30 -(dT)6 and 50 -(dT)10 ssDNA tails in the absence (brown) and presence of the following nucleotides
and analogs: ADP·BeFx (light purple), AMPPNP (yellow ), ADP + Pi (grey), ADP (sky blue), and
ATPγS (sea green). Records were taken at F = 6 pN and smoothed with a Savitzky-Golay window
to 10 Hz (light) and 1 Hz (dark ). (B) Average integrated fluctuations. (C) Integrated fluctuations
at 10 Hz (∆f = 0.07–10 Hz, *P < 0.05, 5 ≤ N ≤ 12). All error bars denote standard error of the
mean.

75

Figure 3.17: Comparison of RecBCD·DNA fluctuations in the presence and absence of ADP·BeFx .
Representative data of the DNA contour length versus time with corresponding histograms fortethers of RecBCD bound to DNA with 30 -(dT)6 and 50 -(dT)10 ssDNA tails in the absence (brown)
and presence of ADP·BeFx (light purple). Records were taken at F = 6 pN and smoothed with a
Savitzky-Golay window to 10 Hz (light) and 1 Hz (dark ).

76

3.6

RecBCD Pin Undergoes Back-and-Forth Translocation Along DNA
Conformational dynamics of the RecBCD·DNA complex in the absence of nucleotides may

be described by three different scenarios to cause back-and-forth changes in the observed DNA
contour length, L (Figure 3.18). In Scenario 1, the enzyme pin does not translocate relative to thessDNA-dsDNA junction, denoted by the first base pair at the junction that remains base paired
when L changes in length by an amount ∆L. Thus, changes in the DNA contour length originate
from conformational changes of the enzyme and ∆xpin = 0, where ∆xpin denotes the change inposition of the RecBCD pin. In Scenario 2, translocation of the RecBCD pin relative to the DNA
substrate is of equal magnitude to that of the observed changes in the DNA contour length, i.e.,∆xpin = ∆L. The enzyme pin translocates by an amount ∆L, destabilizing the equivalent number ofbase pairs. In Scenario 3, a combination of conformational changes of the enzyme and translocationof the enzyme pin relative to the DNA occur such the change in the pin position is nonzero andless than the observed changes of the DNA contour length, i.e., 0 < ∆xpin < ∆L. The pin
destabilizes a portion of the duplex region past the initial junction position, but by a magnitudeless than ∆L. In all three scenarios, when a nucleotide is not bound to the RecBCD, the enzyme
undergoes conformational changes to maintain at least one constant enzyme-DNA interaction thatprevents the enzyme from diffusing backwards along its DNA substrate, a necessary requirement
for unidirectional translocation of a helicase.
To investigate the various mechanisms for the large RecBCD·DNA conformational dynamics
observed, we made two types of modifications to the duplex region upstream of the ssDNA-dsDNA
junction. If the enzyme pin does not undergo translocations relative to the DNA, i.e., if Scenario 1
is true, then the conformational dynamics of the RecBCD·DNA complex should not be affected bychanges made in the duplex DNA region. We modified the sequence of the duplex region (Section
3.6.1) and incorporated an interstrand DNA cross link in the duplex region (Section 3.6.3).

77

Figure 3.18: Potential mechanisms for back-and-forth motion: Three scenarios of RecBCD·DNA
conformational dynamics. The DNA contour length, L, changes in length by an amount ∆L. The
initial position of the ssDNA-dsDNA junction is indicated by the first base pair (pink ). As thejunction position changes by an amount ∆xpin , the newly destabilized base pairs are indicated(pink ). Scenario 1: Conformational changes of the enzyme such that ∆xpin = 0. Scenario 2:
Translocation of the RecBCD pin relative to the DNA substrate such that ∆xpin = ∆L. Scenario 3:
A combination of conformational changes of the enzyme and translocation of the enzyme pin suchthat 0 < ∆xpin < ∆L.

78
3.6.1

Conformational dynamics are dependent on the DNA duplex sequence

We used a set of 50 -(dT)10 -tailed DNA substrates with differing GC content within the first
11 bp upstream of the fork, one with 9% GC content (AT-rich) and one with 91% GC content(GC-rich) (Figure 2.23). The RecBCD·DNA conformational dynamics were markedly different for
RecBCD bound to the AT-rich substrate versus the GC-rich substrate (Figure 3.19), as well asfrom RecBCD bound to a mixed-seqence substrate (Figure 3.20). RecBCD bound to the ATrich construct exhibited conformational dynamics with a mean integrated fluctuations value at
10 Hz of 1.74 ± 0.10 nm (mean ± σ x̄ ), N = 14 (see Appendix C), that was significantly larger(P < 0.04) than that of RecBCD bound to a mixed-sequence construct, which had a mean integrated
fluctuations value at 10 Hz of 1.43 ± 0.09 nm (mean ± σ x̄ ), N = 12 (see Figure 3.15 and Appendix
C). RecBCD bound to the GC-rich construct exhibited conformational dynamics with a mean
integrated fluctuations value at 10 Hz of 1.16 ± 0.07 nm, N = 11 (see Appendix C), that wassignificantly smaller than that of RecBCD bound to the mixed-sequence construct (P < 0.04).
Both AT-rich and GC-rich DNA substrates contained a single-stranded 50 -(dT)10 tail that could
engage the RecD subunit, yet complexes of RecBCD bound to the AT-rich construct exhibited
significantly larger conformational dynamics than complexes of RecBCD bound to the GC-richconstruct. In addition, the conformational dynamics in each case were force-dependent with reduced
conformational dynamics observed with increasing force (see Section 3.6.2).
The sequence-dependence results suggest that the enzyme pin is undergoing translocations
relative to the DNA junction in the absence of ATP, supporting Scenarios 2 and 3 (Figure 3.18).
If RecBCD·DNA interactions were remaining static and the conformational dynamics were solely
caused by enzyme conformational changes, then the dynamics would not depend on the sequenceof the duplex region of the DNA. The sequence-dependent observations suggest that the RecBCD
pin, which has been shown to interact with the ssDNA-dsDNA junction [131], undergoes back-andforth translocations along the DNA substrate in the absence of ATP and that the amplitude of thetranslocations is determined by the GC-content of the DNA duplex region.

79

Figure 3.19: Comparison of RecBCD·DNA fluctuations: RecBCD bound to AT-rich versus GCrich 50 -(dT)10 -tailed DNA. Representative data of the DNA contour length (L) versus time withcorresponding histograms for tethers of RecBCD bound to DNA with 30 -(dT)6 and 50 -(dT)10 ssDNA
tails with GC-rich (pink ) and AT-rich (turquoise) duplex regions. Records were taken at F = 6 pN
and smoothed with a Savitzky-Golay window to 10 Hz (light) and 1 Hz (dark ).

80

Figure 3.20: Sequence-dependence comparison of RecBCD·DNA bound to 50 -(dT)10 -tailed DNA.
(A) Representative data of the DNA contour length versus time with corresponding histograms fortethers of: RecBCD bound to blunt-ended DNA (blue); and RecBCD bound to DNA with a 30 -(dT)6
and 50 -(dT)10 ssDNA tails with GC-rich (pink ), mixed-sequence (brown), and AT-rich (turquoise)
duplex regions. Records were taken at F = 6 pN and smoothed with a Savitzky-Golay window
to 10 Hz (light) and 1 Hz (dark ). (B) Average power spectral density. (C) Average integrated
fluctuations. (D) Integrated fluctuations at 10 Hz (∆f = 0.07–10 Hz, *P < 0.04, 5 ≤ N ≤ 14). Allerror bars denote standard error of the mean.

81
3.6.2

Sequence-dependent dynamics observed at various forces

Force-dependence studies of RecBCD bound to AT-rich, GC-rich, and mixed-sequence 50 (dT)10 -tailed DNA constructs demonstrated the observation of sequence-dependent conformational
dynamics for a range of forces, F = 2, 4, 6, and 10 pN. Three particular complexes of RecBCD
bound to DNA with 30 -(dT)6 and 50 -(dT)10 tails were studied: RecBCD bound to DNA with an
AT-rich base-paired region, a mixed-sequence base-paired region, and a GC-rich base-paired region.
At 2 pN, the mean values (± σ x̄ ) of the integrated fluctuations at 10 Hz were 2.46 ± 0.12 nm,
N = 8 (see Appendix C), 2.66 ± 0.12 nm, N = 7 (see Appendix C), and 3.03 ± 0.47 nm, N = 6 (seeAppendix C) for RecBCD bound to GC-rich, mixed-sequence, and AT-rich DNA, respectively. At
10 Hz, the integrated fluctuations of the GC-rich, mixed-sequence, and AT-rich data were equivalent
within error.
At 4 pN, the mean values (± σ x̄ ) of the integrated fluctuations at 10 Hz were 1.35 ± 0.07 nm,
N = 8 (see Appendix C), 1.61 ± 0.13 nm, N = 6 (see Appendix C), and 2.07 ± 0.21 nm, N = 6(see Appendix C) for RecBCD bound to GC-rich, mixed-sequence, and AT-rich DNA, respectively.
At 10 Hz, the mean value of the integrated fluctuations of the AT-rich data was slightly higher thanthat of the mixed-sequence data (P < 0.10), and that of the mixed-sequence data were slightly
higher than that of the GC-rich data (P < 0.13).
At 6 pN, the mean values (± σ x̄ ) of the integrated fluctuations at 10 Hz were 1.16 ± 0.07
nm, N = 11 (see Appendix C), 1.43 ± 0.09 nm, N = 12 (see Appendix C), and 1.74 ± 0.10 nm,
N = 14 (see Appendix C) for RecBCD bound to GC-rich, mixed-sequence, and AT-rich DNA,
respectively, as discussed in Section 3.6.1. At 10 Hz, the mean value of the integrated fluctuationsof the mixed-sequence data were significantly higher than that of the GC-rich data (P < 0.04), andthat of the AT-rich data was significantly higher than that of the mixed-sequence data (P < 0.04).
At 10 pN, the mean values (± σ x̄ ) of the integrated fluctuations at 10 Hz were 0.88 ± 0.04
nm, N = 8 (see Appendix C), 0.99 ± 0.10 nm, N = 5 (see Appendix C), and 1.12 ± 0.15 nm, N = 7(see Appendix C) for RecBCD bound to GC-rich, mixed-sequence, and AT-rich DNA, respectively.

82
Table 3.1: RecBCD·DNA conformational dynamics of RecBCD bound to 50 -(dT)10 -tailed DNA forvarious duplex sequences.

Force
(pN)
2
4
6
10

DNA
sequence
GC-richMixed
AT-richGC-richMixed
AT-richGC-richMixed
AT-richGC-richMixed
AT-richIntegrated Fluctuations at 10 Hz
mean ± σ x̄ (nm)2.46 ± 0.12
2.66 ± 0.12
3.03 ± 0.47
1.35 ± 0.07
1.61 ± 0.13
2.07 ± 0.21
1.16 ± 0.07
1.43 ± 0.09
1.74 ± 0.10
0.88 ± 0.04
0.99 ± 0.10
1.12 ± 0.15

N
8
7
6
8
6
6
11
12
14
8
5
7

At 10 Hz, the mean values of the integrated fluctuations of the AT-rich and mixed-sequence data
were equivalent within error, and those of the mixed-sequence and GC-rich data were equivalent
within error.
The sequence-dependence data at various forces are summarized in Table 3.6.2, Figure 3.21,
and Figure 3.22. As the applied force was increased, the conformational dynamics of the RecBCD·DNA
complex decreased. Reduced conformational dynamics of RecBCD bound to 50 -(dT)10 -tailed DNA
were observed with increasing force. For each force, RecBCD·DNA conformational dynamics werelargest for the AT-rich DNA, smallest for the GC-rich DNA, and in between the two cases for themixed-sequence DNA. At low applied force, Brownian motion begins to dominate the integrated
fluctuations, which reduces our resolution to measure the sequence dependence of the conformational dynamics of RecBCD·DNA. At higher applied force, the energy required to destabilize theduplex region increases, which reduces the sequence dependence of the conformational dynamics.

83

Figure 3.21: Sequence-dependent conformational dynamics of RecBCD bound to 50 -(dT)10 -tailed
DNA at varying force. Average power spectral density, average integrated fluctuations, and average
integrated fluctuations at 10 Hz (∆f = 0.07–10 Hz) of RecBCD bound to DNA with a 30 -(dT)6
and 50 -(dT)10 ssDNA tails with GC-rich (pink ), mixed-sequence (brown), and AT-rich (turquoise)
duplex regions. (A) Data taken at F = 2 pN. 6 ≤ N ≤ 8 (B) Data taken at F = 4 pN. *P < 0.10,**P < 0.13, 6 ≤ N ≤ 8 (C) Data taken at F = 6 pN. *P < 0.04, 11 ≤ N ≤ 14 (D) Data taken at
F = 10 pN. 5 ≤ N ≤ 8 All error bars denote standard error of the mean.

84

Figure 3.22: Sequence-dependent conformational dynamics of RecBCD bound to 50 -(dT)10 -tailed
DNA versus force. The mean values of the integrated fluctuations at 10 Hz are shown for datataken at F = 2, 4, 6, and 10 pN of RecBCD bound to GC-rich 50 -(dT)10 -tailed DNA (pink ), mixedsequence 50 -(dT)10 -tailed DNA (brown), and AT-rich 50 -(dT)10 -tailed DNA (turquoise). All error
bars denote standard error of the mean.

85
3.6.3

Conformational dynamics were suppressed with an interstrand cross link

To determine whether the RecBCD pin undergoes back-and-forth translocations along DNA,
we measured RecBCD bound to a 30 -(dT)6 and 50 -(dT)10 tailed DNA construct with an engineered,covalent interstrand cross link positioned 3 bp upstream of the junction in the duplex region (“ATcross link, 50 -dT10 ” and “GC cross link, 50 -dT10 ,” Figure 2.23) and compared the DNA contour length
time courses with those of RecBCD bound to a non-cross-linked (dT)-tailed DNA substrate. The
N3 T–butyl–N3 T cross link used bridges a mismatched thymidine base pair while maintaining the
global structure of a non-cross-linked DNA duplex, indicated by circular dichroism measurements[195] and molecular dynamics calculations restrained by nuclear Overhauser effect and dihedral
angle data obtained from NMR spectroscopy [196]. We performed unwinding assays to test theincorporation of the cross link and no unwinding events were observed (data not shown).
In measuring the conformational dynamics of RecBCD complexed with cross-linked DNA, weobserved significantly supressed dynamics (Figure 3.23). The mean integrated fluctuations at 10 Hz
of RecBCD bound to the ends of the cross-linked construct with two AT base pairs prior to the crosslink were 0.96 ± 0.02 nm (N = 7; see Appendix C), 0.47 nm less than that of RecBCD bound to
the ends of a non-cross-linked DNA construct of mixed sequence and 0.12 nm more than RecBCD
bound to blunt-ended DNA. The mean value of the integrated fluctuations at 10 Hz of RecBCD
bound to the ends of the cross-linked construct with two GC base pairs prior to the cross link was
1.02 ± 0.05 nm (N = 5; see Appendix C). The suppressed conformational dynamics indicate that,in the absence of the cross link, the enzyme pin interacts with the duplex region beyond the first 3
bp of the duplex. This evidence for nonzero back-and-forth translocations support Scenarios 2 and3.

3.7

Quantifying RecBCD Destabilization of Duplex DNA
Scenario 2 (∆xpin = ∆L) can be tested with two methods that vary the duplex region of

the DNA to determine at what base pair position the large-scale conformational dynamics of the

86

Figure 3.23: Conformational dynamics are altered with a cross-linked DNA. (A) Representative
data of the DNA contour length versus time with corresponding histograms for tethers of: RecBCD
bound to blunt-ended DNA (blue); RecBCD bound to DNA with 30 -(dT)6 and 50 -(dT)10 ssDNA
tails with an engineered, interstrand cross link 3 bp upstream of the junction, with the first two basepairs comprising of either AT (light blue) of GC (light green) base pairs; and RecBCD bound to
DNA with a 30 -(dT)6 and 50 -(dT)10 ssDNA tails with no cross link (brown). Records were taken at F
= 6 pN and smoothed with a Savitzky-Golay window to 10 Hz (light) and 1 Hz (dark ). (B) Average
integrated fluctuations. (D) Integrated fluctuations at 10 Hz (∆f = 0.07–10 Hz, *P < 0.002,**P < 0.12, 5 ≤ N ≤ 12). All error bars denote standard error of the mean.

87
RecBCD·DNA complex become independent of changes in the duplex region.
In one method, the conformational dynamics of RecBCD bound to 50 -(dT)10 , 30 -(dT)6 tailed
constructs with variations in the sequence of the duplex region are measured. We varied the sequence of the duplex region immediately upstream of the ssDNA-dsDNA junction with the following
sequences (Figure 2.23): 5-bp GC base pairs (“GC-rich”), 3-bp GC base pairs (“Low GC-rich”), 5-bp
AT base pairs (“AT-rich”), and 4-bp AT base pairs (“Low AT-rich”). Because we observe that conformational dynamics of the complex depend on the sequence of the duplex region (Section 3.6.1), we
expect that if the RecBCD·DNA conformational dynamics by the mechanism described in Scenario
2, then the enzyme pin would undergo translocations relative to the DNA with an amplitude equal
to that of the changes in the DNA contour length. The mean values of the integrated fluctuations at
10 Hz for RecBCD bound to the ends of AT-rich and Low AT-rich DNA were 1.74 ± 0.10 nm (mean± σ x̄ ), N = 14, and 1.71 ± 0.19 nm (mean ± σ x̄ ), N = 5, respectively, which were equivalent withinerror. The values of the integrated fluctuations at 10 Hz for RecBCD bound to the ends of GC-richand Low GC-rich DNA were 1.16 ± 0.07 nm (mean ± σ x̄ ), N = 11, and 1.04 ± 0.07 nm (mean± σ x̄ ), N = 5, respectively, which also were equivalent within error. These results suggest that the
enzyme pin does not translocate past the third (fourth) bp when bound to GC-rich (AT-rich) DNA
constructs. Because the peak-to-peak values of the conformational dynamics of the complex weremuch larger than 3–4 bp, this suggests that the RecBCD enzyme undergoes conformational changes,
which contribute to the net conformational dynamics observed of the RecBCD·DNA complex andprovides evidence against Scenario 2. Together with the observations of altered conformational
dynamics in the presence of an interstrand cross link (Section 3.6.3), these results provide support for Scenario 3 (0 < ∆xpin < ∆L) as a mechanism for large conformational dynamics of the
RecBCD·DNA complex.
In a second method, an interstrand cross link can be introduced to examine two critical positions: the position at which the conformational dynamics of the RecBCD·DNA complex begin to be
independent of cross-link position and the position at which the cross link inhibits pin translocation
to zero (∆xpin = 0). If the enzyme pin does not translocate beyond the third or forth base pair,

88

Figure 3.24: (A) Representative data of the DNA contour length versus time with corresponding
histograms for tethers of: RecBCD bound to Low GC-rich DNA (green), GC-rich DNA (pink ), Low
AT-rich DNA (orange), and AT-rich DNA (blue) Records were taken at F = 6 pN and smoothedwith a Savitzky-Golay window to 10 Hz (light) and 1 Hz (dark ). (B) Average power spectral
density. (C) Average integrated fluctuations. (D) Integrated fluctuations at 10 Hz (∆f = 0.07–10
Hz, *P < 0.05, 5 ≤ N ≤ 14). All error bars denote standard error of the mean.

89
as suggested in the AT-rich (GC-rich) vs Low AT-rich (Low GC-rich) sequence-dependent studies, then the conformational dynamics of the RecBCD·DNA complex would be equivalent in thepresence and absence of an interstrand cross link positioned at the fifth base pair. If instead the
back-and-forth pin translocation were completely inhibited by positioning an interstrand cross linkat the first base pair at the junction, then the DNA contour length records could be analyzed to
examine the conformational dynamics of the enzyme. These two studies have yet to be performed.
We measured the conformational dynamics of an interstrand cross link at the third base pairfrom the ssDNA-dsDNA junction (Section 3.6.3). At this position, the RecBCD pin is able totranslocate at most by 2 bp (0 ≤ ∆xpin ≤ 2 bp, or 0 ≤ ∆xpin ≤ 0.68 nm). Interestingly, the L timecourses of RecBCD bound to the cross-linked DNA exhibit distinct, “spiked” features that contrast
with the L time courses of RecBCD bound to blunt-ended DNA (Figure 3.25). The number of
0.87-nm spikes (see Section 2.7) within conformational dynamics records of RecBCD bound to a50 -(dT)10 -tailed, AT cross-linked DNA was 45.9 ± 2.8 (mean ± σ x̄ ), N = 7 (Figure 3.25). This was
significantly larger than the number of spikes observed in records of RecBCD bound to blunt-ended
DNA (P < 0.0007; see Section 2.7), which were 17.0 ± 4.2 (mean ± σ x̄ ), N = 5. The number of
0.87-nm spikes within conformational dynamics records of RecBCD bound to a 50 -(dT)10 -tailed, GC
cross-linked DNA was 50.0 ± 3.8 (mean ± σ x̄ ), N = 5, which was also significantly larger than thenumber of spikes observed in records of RecBCD bound to blunt-ended DNA (P < 0.0005) (datanot shown). Also, almost no spikes were detected in the records of DNA directly attached to thesurface, which were 2.2 ± 0.7 (mean ± σ x̄ ), N = 10, and were significantly less than the numberof spikes observed in RecBCD bound to blunt-ended DNA (P < 0.03). The spikes exhibited by
RecBCD bound to cross-linked 50 -(dT)10 -tailed DNA may be due to the interactions of RecBCD
encountering the cross link and/or due to conformational dynamics of RecBCD. When RecBCD
is bound to the ends of blunt-ended DNA, the RecD motor is not engaged by the 50 DNA, and
RecBCD does not assume a dynamically fluctuating state.

90

Figure 3.25: Comparison of number of spikes in RecBCD bound to cross-linked versus non-crosslinked DNA. (A) Representative records of the DNA contour length of RecBCD bound to bluntended DNA (blue) and RecBCD bound to an AT cross-linked DNA with 30 -(dT)6 and 50 -(dT)10
ssDNA tails (light blue), with corresponding histograms. Data were taken at F = 6 pN and smoothedwith a Savitzky-Golay window to 10 Hz (light) and 1 Hz (dark ). (B) The average number of 0.87-nm
spikes observed in the 1-Hz DNA contour length records of DNA tethered to the surface (black ),RecBCD bound to blunt-ended DNA (blue), and RecBCD bound to an AT cross-linked DNA with30 -(dT)6 and 50 -(dT)10 ssDNA tails (light blue). *P < 0.0007, **P < 0.03, 5 ≤ N ≤ 10. Error bars
denote standard error of the mean.

91

3.8

Discussion

3.8.1

Large conformational dynamics present in the absence of ATP

With a high-precision optical-trapping microscope, we observed multi-base-pair conformational dynamics of RecBCD complexed with a DNA substrate in the absence of ATP. This indicatesthat the large, multi-base-pair motions are not powered by ATP hydrolysis or binding, but must
be strictly due to interactions between the enzyme and the DNA substrate. Specifically, our observations suggest that the conformational dynamics are dependent on interactions between the RecD
motor and the 50 -ssDNA end. The conformational dynamics may be driven by RecD translocations
relative to the 50 ssDNA end, or by RecD-DNA interactions that allosterically cause conformationalchanges elsewhere in the holoenzyme. Interestingly, our records indicate that only a 50 -(dT)10 ssDNA tail is required for the 50 DNA to interact with the RecD motor, which is consistent withequilibrium binding studies that indicate a plateau in the equilibrium binding constant of RecBCD
bound to the ends of DNA duplexes with 50 -(dT)n ssDNA tails for values of n ≥ 10 [190]. Ina crystal structure of RecBCD bound to a single-stranded 50 tail comprised of seven 5-iodouracil
nucleotide analogs and three nucleotides, comprising a 50 tail of 10 repeats, the 50 tail appears to
only partially engage the motor domains of the RecD motor [191]. Our measurements indicate that
this high-resolution snapshot of RecBCD·DNA in a crystal lattice must be only one frame in a dynamic motion picture of both rapidly-fluctuating, back-and-forth translocation of the RecBCD pin
along the DNA and conformational dynamics of the enzyme, which contribute to the RecBCD·DNA
conformational dynamics that are on the scale of multiple base pairs.
RecBCD does not directionally translocate when paused on an unwound DNA substrate,suggesting that a part of the enzyme maintains constant contact with a part of the DNA substrate,either in the 30 single-stranded region, the 50 single-stranded region, the duplex region, or somecombination. Crystal structures show that RecBCD extends an “arm” which contacts the duplex
region 12 bp beyond the junction [131, 191], which is one potential interaction that may remain
constant during the conformational dynamics.

92
3.8.2

RecBCD operates by dynamic destabilization and conformational changes ofthe enzyme

Notably, the distinctive alteration of the conformational dynamics when RecBCD is boundto a cross-linked DNA substrate indicates that the RecBCD pin translocates back and forth along
the DNA in the absence of ATP. The large conformational dynamics cannot simply be attributed
solely to conformational changes of the enzyme with positioning of the enzyme staying relatively
constant to that of the DNA substrate. Our observations are consistent with the part of RecBCD
that contacts the junction, the RecC pin, undergoing translocation relative to the DNA and thereby
contributing to active destabilization of the duplex. Additionally, the large conformational dynamics
cannot be attributed to translocations of the enzyme pin of equal magnitude. Our observations are
consistent with RecBCD pin translocation relative to the DNA of 3–4 bp, which are smaller thanthe peak-to-peak behavior of the complex, suggesting that conformational changes of RecBCD
contribute to the net conformational dynamics of the RecBCD·DNA complex.
The conformational dynamics are a direct, real-time observation that include dynamic destabilization of the double-stranded region of the DNA substrate by the RecBCD helicase. The directdestabilization of dsDNA by an active helicase to promote DNA opening has been modeled [110].
Correlation of the conformational dynamics with unwinding initiation [32] suggests the conformational dynamics are an integral aspect to the mechanism of the DNA unwinding mechanism of
RecBCD. Because RecBCD has the ability to unwind 6 bp in the absence of ATP [189, 190, 191],
it is not contradictory that RecBCD is able to dynamically destabilize multiple base pairs at the
DNA junction in the absence of ATP. The large conformational dynamics could function to makethe cleft between the motor domains more accessible for fast ATP binding and ADP + Pi release,
contributing to the high unwinding velocities achieved by this enzyme .
The reduction of the conformational dynamics of RecBCD·DNA in the presence of ADP·BeFx ,AMPPNP, or ADP + Pi suggests that the RecBCD·DNA complex tightens down on the DNA when
ATP binds to RecBCD. Presumably, the ATP analog binds to either the RecB or RecD ATP-binding

93
clefts, causing a conformational change in the RecBCD·DNA complex that reduces the large, multibase-pair conformational dynamics. Structural studies of a RecD homolog, RecD2 from Deinococcus
radiodurans, determined 2.5-Å resolution structures of RecD2 complexed with ssDNA in the presence
and absence of an ATP analog [28]. Our observation of reduced conformational dynamics in thepresence of ADP·BeFx and other nonhydrolyzable ATP analogs is consistent with this structural
result, which indicates that upon ATP-analog binding the number of contacts between RecD2 andthe DNA substrate increase and the cleft between the 1A and 2A motor domains closes, tightening
the interaction between enzyme and DNA.
Together, our observations suggest a RecD motor mechanism that includes dynamic destabilization, in which conformational dynamics of the RecBCD·DNA complex are modulated by the
nucleotide-bound state of the enzyme. When no ATP is bound and RecD is engaged by the 50
ssDNA end, conformational changes in the RecBCD·DNA complex occur, resulting in conformational changes in the RecBCD and active destabilization of the ssDNA-dsDNA junction, which isconsistent with back-and-forth translocation of the dual-methionine RecC pin beyond the averagejunction position by multiple base pairs. A mechanism that includes dynamic destabilization may
aid in RecBCD’s fast unwinding speed as well as its ability to displace DNA-bound proteins. It
is remarkable that a rearrangement of the three SF1 subunits, RecB, RecC, and RecD, into one
holoenzyme can create such a robust and multifunctional enzyme.

Chapter 4

glmS Ribozyme Cleavage

4.1

Active Roles of RNA

4.1.1

Ribozymes are catalytic RNA molecules

Molecular biologists have long suspected that RNA molecules played a key role in the processof how life began on Earth. RNA has an essential role in protein synthesis, a fundamental process
within all cells. One of the first observations of RNA catalytic activity in the absence of proteins wasof the self-splicing pre-ribosomal RNA of the ciliate Tetrahymena [34]. With this work, the word
“ribozyme” was coined for the general concept of an RNA molecule with enzymatic activity. The
first example of a ribozyme that exhibited multiple turnover was the RNA subunit of ribonuclease
P (RNase P), which hydrolyzes the 50 -end of precursor transfer RNA (tRNA) [33]. These findings
provided further credibility to the hypothesis of an RNA world [197].
In the RNA world hypothesis, RNA served both as the genetic material and the principal
cellular enzyme in cells [198]. This hypothesis postulates that as cellular metabolism became moresophisticated, increasing demands on biocatalysts provided the need for the transition to protein
enzymes. Naturally-occurring ribozymes provide further evidence for the hypothesized RNA world[197]. Such ribozymes are present in organisms ranging from bacteria to humans. Examples include RNA that performs self-cleavage via transesterification (e.g., hammerhead, hairpin, hepatitis
delta virus, and Varkud satellite ribozymes); self-splicing via transesterification (e.g., Group I intron and Group II intron); pre-tRNA processing via hydrolysis (e.g., RNase P); RNA splicing via

95transesterification (e.g., spliceosome); and peptidyl transfer (e.g., ribosome).
In vitro selection experiments demonstrate that it is possible for functional nucleic acids toarise from random sequence libraries [199, 200]. This technique was coined the systematic evolutionof ligands by exponential enrichment (SELEX) [201, 202]. Random sequence DNA or RNA pools
can be generated by a combination of chemical and enzymatic synthesis, and molecules that performa particular function can be selected from the pools. For example, the molecules might be selected
to exhibit particular catalytic functions, i.e., ribozymes, or to exhibit particular binding properties,i.e., RNA “aptamers,” structured nucleic acids capable of specifically binding a target molecule [203].
The selected species are then amplified. After multiple cycles of selection and amplification to mimic
natural selection, the RNA molecules generated are typically highly-fit molecules. For example,
the theophylline aptamer binds theophylline with a Kd of ∼400 nM and can discriminate against
caffeine, which differs by a single additional methyl group, by a factor of 10,000 [204]. These selectednucleic-acid molecules have been evolved to contain specific binding pockets, perform catalysis, bindcofactors, and adopt complex three-dimensional structures [200].

4.1.2

Riboswitches detect specific metabolites

A riboswitch is a structured RNA element that modulates gene expression in response to thedirect binding of a metabolite [205]. Riboswitches, which usually reside in the 50 noncoding regionsof messenger RNA (mRNA), are cis-acting genetic control modules, meaning they act on the mRNAin which they are located. They are generally utilized in the feedback repression of genes involvedin the synthesis or import of the cognate metabolite. To control gene expression, riboswitches form
highly-specific binding pockets for their target metabolite and typically undergo allosteric changes
in structure upon ligand binding (Figure 4.1). This interaction of the aptamer domain with theimmediately downstream functional domain, the “expression platform,” provides the interface withvarious other components of the gene-expression apparatus. Ligand binding is achieved solely by
the aptamer domain, which usually does not require the presence of the expression platform or theaction of protein factors [205]. Over 20 distinct classes of riboswitches have been identified [206].

96

Figure 4.1: Example of gene expression regulation by a riboswitch: Riboswitch-regulated transcription termination. Reprinted from Garst and Batey (2009) [207], with permission from Elsevier.
The aptamer domain (black /gray) forms prior to transcription of the expression platform (cyan),allowing this receptor to bind its metabolite (M; orange). Metabolite binding collapses the aptamer
domain into a conformation that sequesters the switching sequence (SS) from pairing with the expression platform, leading to formation of a terminator stem (T), which attenuates transcription.
In the event that ligand binding does not occur, particular elements of the aptamer domain remainloosely structured, leading to formation the antiterminator stem (AT), allowing RNA polymerase(RNAP) to continue transcribing.

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Figure 4.2: Reaction catalyzed by glucosamine-6-phosphate synthase. Reprinted from Milewski,
Copyright (2002) [212], with permission from Elsevier.

Riboswitches are usually adjacent to genes involved in the synthesis or transport of their
metabolite ligand [205]. Many candidates have been identified in searches for highly-conserved
RNA domains that reside in intergenic regions of bacterial genomes [208, 209, 210]. Thus, thediscovery of riboswitches has served as a powerful tool for inferring functions of uncharacterized
genes [208, 211].

4.1.3

The glmS ribozyme-riboswitch

The glmS ribozyme was the first natural catalytic RNA discovered to require a small-molecule
activator for catalysis [54]. This ribozyme-riboswitch is a conserved element in numerous Grampositive bacteria in the 50 untranslated region (UTR) of the mRNA encoding glucosamine-6-phosphate(GlcN6P) synthase. It functions by undergoing ligand-induced, site-specific self-cleavage that reduces gene expression through destabilization of the mRNA [54, 55]. GlcN6P synthase, formallyknown as glutamine-fructose-6-phosphate amidotransferase, is an enzyme that uses fructose-6phosphate and glutamine to generate GlcN6P [212] (Figure 4.2). GlcN6P is the ligand that binds
to the glmS ribozyme, providing a negative feedback signal for synthesis of the enzyme that produced it. The generation of GlcN6P is the first step in the pathway for the production of uridine
50 -diphospho-N -acetyl-D-glucosamine, which is subsequently used in the process of cell-wall biosynthesis.
X-ray crystallography studies reveal the architecture of the glmS ribozyme [213, 214, 215, 216].
The crystal structure of the glmS ribozyme with GlcN6P bound in the active site was solved to 2.5-

98
Å resolution [214] (Figure 4.3). The glmS ribozyme is ∼50 Å wide and ∼20 Å deep. It is comprisedof a short, helical segment surrounded by two sets of coaxially-stacked helices . The short, central
stack, P2.1, is sandwiched by a long stack, consisting of P1 and P3.1, and a smaller stack, consistingof P4 and P4.1.
The ribozyme core consists of a double-pseudoknot structure that comprises both the active
site and the metabolite-binding pocket. The positioning of the central helix P2.1 and the adjacenthelix P2.2, as well as that of a tightly bound metal ion, form the double pseudoknot by four
nonhelical crossovers that connect the P1-P2.2-P2 stack with P2.1.
The ribozyme peripheral, comprised of P3, P3.1, P4, and P4.1, has been shown to be necessary
for optimal self-cleavage rates at physiological Mg2+ concentrations [217, 218]. This accessory
domain is anchored in position by a third pseudoknot between P3 and P3.1 at one end and by atertiary interaction between P1 and and P4.1 at the other (Figure 4.3). Extensive contacts between
the A-rich internal loop of the P4 element and the P2.1 minor groove serve to further buttress thecatalytic core.
The atomic-resolution structures of the free and ligand-bound forms of the glmS ribozyme
are nearly superimposable, indicating that ligand binding does not induce a conformational change
in the RNA, as is typical of other riboswitches [213, 214, 215, 216].
In addition to the observed crystalline states of the glmS ribozyme, solution studies provide
further evidence for a prefolded active site within a ribozyme structure that does not undergo a global
conformational change upon GlcN6P binding [54, 219, 220]. Fluorescence resonance energy transfer(FRET) studies show that the global conformation does not change upon ligand binding [220].
Cross-link studies demonstrate that a UV cross-linked complex still exhibits catalytic activity [219].
Collectively, the crystallographic, FRET, and cross-link studies, indicating a rigid, preorganized fold,suggest that GlcN6P functions not as an allosteric effector but as a coenzyme, directly participatingin catalysis.
Since the discovery of the glmS ribozyme, there has been evidence for a eukaryotic ribozyme
riboswitch [221]. The CLEC2 ribozymes, variants of the hammerhead ribozyme found as a dis-99

Figure 4.3: Structure of the glmS ribozyme. Reprinted from Cochrane et al., Copyright (2007) [214],with permission from Elsevier. (A) Secondary structure of the Bacillus anthracis glmS ribozyme.
(B) Tertiary structure depicted in ribbons format of the P1 helix (red ); P2 helix (yellow ); P3–P4
helices (green); double pseudoknot consisting of helices P2.1, P2.2, P2a, and related joiner regions(blue); and U1A and the U1A binding loop (gray). GlcN6P is bound in the active site and is shownin ball and stick representation.

100
continuous sequence in the 30 UTR of rodent C-type lectin type II (clec2) genes, can self-cleaveand reduce clec2 gene expression in mouse cells. Although the specific effector for this putative
riboswitch is still unknown, a mechanism controlling cleavage seems likely because if the ribozyme
always cleaved, the clec2 genes would never be expressed. The discovery of the glmS and CLEC2
ribozymes constitute specific examples of what may be a far more general regulatory phenomenon[222].
The study of the glmS ribozyme as a naturally-occurring biosensor may advance the development of artificial RNA-based biosensors. Aptamers, as discussed in subsection 4.1.1, aresingle-stranded nucleic acid molecules that bind their cognate ligands with high binding affinity
and specificity. In this sense, they are comparable to monoclonal protein antibodies [223]. The
development of aptamers has been accompanied by a growing range of applications in biosensor andmedical technologies [53, 224].
In comparison with monoclonal antibodies, which are produced in cells, aptamer biosensors
carry some distinct advantages [53]. First, aptamer biosensors can be selected for toxic and nonimmunogenic targets. Second, modifications to the biosensor and assay conditions can be easily
customized for the immobilization of biochips. Third, aptamers prove to be more resistant todenaturation than antibodies [225]. Fourth, aptamers have been shown to be active in the presenceof organic solvents [226]. Finally, aptamers can be produced on a large scale though polymerase
chain reaction and in vitro transcription, which is highly reproducible and inexpensive.
In this context, study of the glmS ribozyme provides insight into a mechanism for gene regulation by a ribozyme, as well as an example of an RNA-based biosensor, which may be applicableto further developments within the field of biosensors. Here, a single-molecule cleavage assay of theglmS ribozyme is developed and used to measure cleavage of single ribozymes. This proof-of-concept
study demonstrates an assay for investigating nucleic-acid-based biosensors.

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Figure 4.4: Schematic of the single-ribozyme cleavage assay. A RNA:DNA hybrid molecule is held
between two points, a cover slip and a nanoparticle. The ribozyme (red ) and DNA handles (black )are shown. Upon binding of glucosamine-6-phosphate (GlcN6P), the glmS ribozyme undergoes
self-cleavage.

4.2

Materials and Methods
To allow monitoring of single cleavage events, single glmS ribozymes were immobilized ona surface. Each ribozyme was attached to a surface and to a 180-nm diameter nanoparticle via
RNA:DNA hybrid handles and biological linkages (Figure 4.4). In our experiments, polystyrene
nanospheres were antidigoxigenin-coated and adhered to the 30 end of the digoxigenin-labeled DNA
bound to the 50 RNA terminus. The 50 end of the DNA bound to the 30 terminus of the RNA
was biotinylated and bound tightly to the glass substrate via a streptavidin and biotinylated-BSA
molecule. Upon addition of GlcN6P molecules, the ribozyme underwent self-cleavage and quickly
diffused away from the surface.
We used two strategies to maintain attachment of each nanoparticle to the surface by a single
RNA:DNA construct. First, we designed an RNA:DNA construct with very short handles of 30 basepairs each. This length is short compared to the diameter of the nanoparticle and spatially hinders
simultaneous attachment of multiple RNA:DNA constructs to the same nanoparticle and to thesurface. Second, when incubating nanoparticles with the RNA:DNA construct, we incubated witha 5:1 excess of nanoparticles, limiting the number of nanoparticles with more than one attachment.
As confirmation that we do not observe kinetics due to multiply-tethered nanoparticles, we verified
that the cleavage data do not fit to multiply-tethered nanoparticles by fitting the data to y(t) =y0 + B1 exp [−k(t − t0 )] + t B2 exp [−k(t − t0 )], where y is the fraction cleaved, t is the time after

102
GlcN6P is introduced, y0 is an offset in y, B1 and B2 are amplitude coefficients, and k is the pseudorate of cleavage.

4.2.1

RNA:DNA hybrid constructs

To tether single molecules of RNA to a nanoparticle and a surface, we used biotinylatedDNA and digoxigenin-labeled DNA at the ends of RNA:DNA constructs. Using hybrid constructs
ensures that each tether will be hybridized on both sides of the ribozyme. Standard polymerase
chain reaction (PCR) and cloning strategies were used to create a DNA plasmid comprising a T7
RNA polymerase promoter, a sequence complementary to the Handle A DNA, a 161-nucleotide
insert of the glmS ribozyme, and a sequence complementary to the Handle B DNA (Figure 4.5).
It has been shown that glmS ribozymes possess considerable self-cleavage activity in theabsence of ligand in TRIS-buffered solution [54, 227]. Thus, most reactions with the glmS RNA
were performed in HEPES buffer. The glmS RNA was RNA was transcribed in vitro from PCR
products. Transcription reactions consisted of 30 mM HEPES (pH 8.0), 10 mM dithiolthreitol,
0.1% Triton X-100, 0.1 mM spermidine-HCl, 4 mM each nucleotide triphosphate (Sigma-Aldrich,
St. Louis, MO; pH adjusted to 8.0), 32 mM MgCl2 , 50 µg/ml T7 bacteriophage RNA polymerase,
1 unit/ml inorganic pyrophosphatase (Sigma-Aldrich), and template DNA at 9 µg/ml. Reactions
were incubated for 2 h at 37 ◦ C. The transcribed product was heated for 1 min at 90 ◦ C andimmediately ran on a denaturing polyacrylamide gel (8%; in TBE buffer with urea). The RNA waseluted overnight in an elution trap at 4 ◦ C in TBE, then purified in an exchange column in 50 mM
potassium HEPES (pH 7.5) and centrifuged at 36.5 krpm.
Short, 30-nt ssDNA oligomers, denoted “Handle A” and “Handle B,” were end-labeled witheither biotin or digoxigenin (Integrated DNA Technologies, Coralville, IA). Handle A and Handle
B DNA are complementary to the 50 and 30 end sequences of the RNA, respectively. DNA andRNA were annealed with a DNA:RNA ratio of 4:1 and incubated for 80 min on ice. The annealed
and unannealed DNA were separated using a nondenaturing polyacrylamide gel and recovered using
standard crush-and-soak methods.

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Figure 4.5: Sequence of the glmS -ribozyme-containing RNA used in the RNA:DNA construct. A
233-nt RNA construct was designed to contain the glmS ribozyme (blue) flanked with handles thatbase pair with DNA Handle A (red ) and DNA Handle B (green). The RNA sequence (top) is shown.
The cleavage site, indicated in bolded letters, is located near the 50 -end of the glmS ribozyme region.
The DNA template for the RNA (bottom) is shown. Restriction-enzyme sites are underlined.

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4.2.2

Protein-conjugated beads

To make the antidigoxigenin-coated nanoparticles, we coupled antidigoxigenin fragment-antigenbinding (Fab) fragments (Roche Applied Science, Indianopolis, IN) to 0.18-µm diameter spherical
nanoparticles (Aldehyde/Sulfate latex particles; Interfacial Dynamics, Portland, OR; now Invitrogen, Carlsbad, CA) [87]. First, we passivated two 1.5 ml plastic tubes for 2 h at room temperature
with 1.2 ml 2 mg/ml acetylated BSA (acBSA; Sigma-Aldrich) in phosphate buffered saline (PBS).
The tubes were then rinsed with RNase-free (DEPC-treated) H2 O. One tube was used during theprotocol and the other was used to store the final modified antidoxigenin beads at 4 ◦ C.
Next, in the protocol tube, we washed latex beads (100 µl of 4.4% w/v 180-nm beads) 4 times
in 1 ml PBS buffer (centrifuge for 9 min at 14 krpm to form a pellet, remove supernatant, resuspend
in 1 ml PBS and pipette and vortex to suspend). After washing, beads were resuspended in 500µl PBS with 20 µl of 80 mg/ml acBSA. Beads were sonicated for 40 min, or until fully separated.
Next, we added 500 µl of 2 mg/ml anti-digoxigenin Fab fragments in PBS to the beads, mixed by
pipetting, and incubated on a rotator (> 24 h). After incubation, we centrifuged the beads to forma pellet (8 min at 14 krpm), removed the supernatant, and re-suspended them in 1.2 ml of 1.0 M
glycine in PBS, incubating for 40 min on a rotator at room temperature to quench the reaction.
Next we centrifuged again to form a pellet (8 min at 14 krpm) and removed the supernatant. We
performed the final resuspension in PBS, 2.9 mg/ml acBSA, and 2% (w/v) sodium-azide, to form
the final solution (∼2 nM for 180 nm beads). The final, modified-bead solution was transferred tothe other acBSA-passivated tube.

4.2.3

RNA tethered-bead assay with BSA surface passivation

One method to passivate the flow chambers was to use bovine serum albumin (BSA) and biotinylated BSA (bio-BSA) to immobilize the nucleic-acid constructs while rejecting protein adsorption (see Appendix A). BSA served to passivate the surface to prevent nonspecific attachment of theanti-digoxigenin-coated nanoparticles. Biotinylated BSA allowed specific attachment of RNA:DNA

105
constructs through a biotin-streptavidin linkage. To passivate the cover glass surface, a cover glass(20 x 40 mm, Corning Life Sciences) was first cleaned with potassium hydroxide dissolved in denatured ethanol. Next, flow chambers with an internal volume of 15 µl were constructed from acleaned glass cover slip, a standard microscope slide (Corning Life Sciences), double-stick tape (3M,
St. Paul, MN), and 5-minute epoxy (Devcon, Danvers, MA). The flow chamber was then incubatedwith a 125 µg/ml bio-BSA in 0.1 M sodium phosphate buffer for 20 min, washed with binding buffer[10 mM HEPES (pH 7.5), 150 mM potassium chloride, 0.4% Tween-20, and 3 mg/ml acetylated
BSA] and incubated for 10 min.
Flow cells were next incubated with streptavidin (10 µg/ml in binding buffer) for 10 min,then washed with 400 µl of binding buffer. Nanoparticles were sonicated, then incubated 1 hwith RNA:DNA constructs at a 5:1 mol ratio (900 pM nanoparticles) in binding buffer [10 mM
HEPES (pH 7.5), 150 mM KCl, 0.4% Tween-20 (BioRad, Hercules, CA), and 3 mg/ml acBSA].
Next, nanoparticle-oligo complexes were flowed through the flow cells at a concentration of 188 pM
nanoparticles. After 20 min of incubation, flow cells were washed with 400 µl of binding buffer. The
enzymatic reaction was initiated by flowing in 50 µl reaction buffer [50 mM HEPES (pH 7.5), 150
mM KCl, 1 mM MgCl2 , and varying concentrations of GlcN6P].

4.2.4

RNA tethered-bead assay with PEG surface passivation

A second method to tether RNA to passivated flow chambers utilized polyethylene glycol(PEG) surfaces [228]. We used customized protocols for covalently attaching PEG molecules to
the glass slide and cover glass surface, which involved careful storage and handling of the materials,
details of which are explained in [87]. The protocol is briefly summarized here. First, slides and coverglasses were plasma-cleaned, then KOH-cleaned. Next, we performed aminosilanization on the glass
surfaces by incubating for 20 min in a solution containing aminopropylsilane [N-(2-Aminoethyl)3-Aminopropyltrimethoxysilane (UCT A0700)] and methanol. After aminosilanization, we rinsed
with methanol, diH2 O, 1M KOH, and plenty of diH2 O, and dried with nitrogen gas. We thenassembled the aminosilanated cover glass and glass slide into flow cells with no epoxy.

106
Next, PEG was covalently attached using a 60:1 ratio mix of methoxy-PEG (mPEG) andbiotinylated-PEG (bio-PEG) molecules, dissolved in 0.1 M sodium bicarbonate bu?er. PEG solution
was flowed into the assembled, aminosilanated samples, which were then incubated for 3–4 hours ina dark humidity chamber. After incubation, we rinsed the ßow cells with diH2 O, dried them withnitrogen, and stored them in black-tape wrapped bottles that were vacuum sealed.
Flow cells were rinsed with 200 µl of Tris buffered saline [TBS; 20 mM Tris-Cl (pH 7.3) and200 µl of 150 mM NaCl] and Wash Buffer [WB; 10 mM HEPES (pH 7.3), 100 mM KCl, and 1 mM
MgCl2 ] before incubated with streptavidin (0.1 mg/ml in TBS) for 30 min, then washed with 400µl of TBS and 400 µl of WB. Nanoparticles were sonicated, then mixed and incubated 1 h with
RNA:DNA constructs at a 5:1 mol ratio (375 pM nanoparticles) in binding buffer [10 mM HEPES(pH 7.5), 50 mM KCl, and 3 mg/ml filtered acBSA] at 4 ◦ C. Next, nanoparticle-oligo complexes
were flowed through the flow cells at a concentration of 188 pM nanoparticles and incubated for 20
min at 4 ◦ C. Then, flow cells were washed with 600 µl of WB. The enzymatic reaction was initiatedby flowing in 50 µl reaction buffer [50 mM HEPES (pH 7.5), 150 mM KCl, 1 mM MgCl2 , andvarying concentrations of GlcN6P].

4.2.5

Single-molecule data acquisition and analysis

We used tethered-particle microscopy and measured single cleavage events in real time. A
wide-field differential-interference-contrast (DIC) inverted microscope (TE 2000-S; Nikon Corporation, Japan) with 100x oil immersion objective (Nikon Plan Fluor, 100x/1.30 oil DIC H; Nikon
Corporation, Japan) was used to image an area of 100 x 100 µm2 to a charge-coupled device camera(CCD100; Dage-MTI, Michigan City, IN). Cleavage events were observed as previously immobile
nanoparticles that suddenly diffused away from the cover-glass surface.
The recorded image was digitized and analyzed (Vision Assistant; National Instruments,
Austin, TX). Analysis included enhancing the brightness and contrast of the image, converting
the image to binary, calculating the sum signal of the region of interest for each nanoparticle,
and determining in which frame the nanoparticle disappeared from the image with a customized,

107

Figure 4.6: Signal of a cleavage event as measure with a tethered-particle assay. The time course isa representative trace of the enhanced sum signal measured by the CCD camera of a selected regionof interest. An automated algorithm determines the start time (blue square) and the cleavage time
(red circle).

automated procedure (Igor Pro; WaveMetrics, Portland, OR) (Figure 4.6). The threshold value wasset halfway between the maximum intensity of the region of interest and the background.

4.2.6

Ensemble FRET experiments

We also studied freely diffusing RNA:DNA constructs using fluorescence resonance energy
transfer (FRET). Here, we used RNA:DNA constructs with a 34-nt DNA labeled with Alexa-546
at the 50 end annealed to the 50 end of the RNA and 41-nt DNA labeled with Alexa-647 at the
30 end annealed to the 30 end of the RNA. Fluorescence values were measured using a fluorometer(Fluorolog-3™; Jobin Yvon, now HORIBA Jobin Yvon Inc, Edison, NJ), FluorEssence™ software,and a 50 µl cuvette. The FRET signal was measured by exciting the sample at 530 nm, whichprimarily excites the donor, and collecting the acceptor emission at 667 nm, with excitation andemission slits set to a 5-nm bandpass. Annealed ribozymes (final concentration 5 nM) were incubatedat 25 ◦ C for at least 15 min in 50 mM HEPES (pH 7.5) and 150 mM KCl, and then transferred to a
50 µl cuvette. GlcN6P and MgCl2 was manually added to concentrations of 30 umuM and 1 mM,
respectively. Photobleaching was negligible, as indicated by measurement of the acceptor emission
spectrum before and after cleavage (data not shown).
The decrease in FRET efficiency from cleavage resulted in a time-dependent decrease in

108
acceptor emission. These data were used to determine the apparent rate constant for cleavage, kobs ,by fitting the FRET acceptor fluorescence, A, to a single exponential,
A(t) = A0 e−kobs t + Aoffset ,(4.1)

where kobs is the observed rate of cleavage, A0 is the fluorescence at time zero, and Aoffset is thechange in fluorescence between time zero and infinite time. The normalized acceptor fluorescence
was normalized by the equation y = (A − Aoffset )/A0 .

4.3

Preliminary Results

4.3.1

Cleavage of single glmS ribozymes

We observe cleavage events of the surface-immobilized ribozyme by video microscopy. Enzymatic activities of the ribozymes were monitored by observing when the nanoparticle left thesurface. The reaction time course is shown in Figure 4.7. The fraction cleaved was fit to the double
exponential equationy(t) = y0 + C1 exp (−k1 t) + C2 exp(−k2 t),(4.2)

where y is the fraction cleaved, t is the time after GlcN6P is introduced, y0 is the offset in y, C1 andC2 are amplitude coefficients, and k1 and k2 are the pseudo-rate constants of cleavage. The data
do not fit to a single exponential, which suggests heterogeneous reaction kinetics that are describedwith two or more distinct cleavage-rate constants. We observed cleavage rates from one flow cell of
k1 = 1.96 ± 0.30 min−1 and k2 = 0.28 ± 0.13 min−1 and amplitude coefficients of C1 = −0.67 ± 0.09
and C2 = −0.33 ± 0.03 at higher concentration of ligand [50 mM HEPES (pH 7.5), 150 mM KCl, 1
mM MgCl2 , and 200 µM GlcN6P] on a BSA-coated surface. We also observed slower cleavage rates
from the combined data of two flow cells of k1 = 0.82 ± 0.07 min−1 and k2 = 0.06 ± 0.04 min−1
and amplitude coefficients of C1 = −0.72 ± 0.04 and C2 = −0.35 ± 0.07 at lower concentrationof ligand [50 mM HEPES (pH 7.5), 150 mM KCl, 10 mM MgCl2 , and 50 µM GlcN6P] on PEGcoated surfaces. Preliminary results indicate double-exponential kinetics that were present on both
BSA-passivated and PEG-passivated surfaces.

109

Figure 4.7: Single-molecule measurements of glmS ribozyme cleavage. The blue open circles show
the reaction time course of the surface-immobilized ribozyme. The dashed black (solid green) curve
is a numerical fit to a single (double) exponential. (A) Ligand is present. Measurement of one flow
cell of glmS ribozymes in in 50 mM HEPES (pH 7.5), 150 mM KCl, 1 mM MgCl2 , and 200 µM
GlcN6P. (B) No ligand is present. Measurement of one flow cell of glmS ribozymes in in 50 mM
HEPES (pH 7.5), 150 mM KCl, and 1 mM MgCl2 .

110

Figure 4.8: Real-time ensemble measurement of glmS ribozyme cleavage. Time dependence of theacceptor fluorescence after mixing with GlcN6P and Mg2+ . Rates of cleavage were measured (bluedots) and fit to a single exponential (black line).

We also performed control experiments to test whether glmS ribozymes were undergoing selfcleavage in the absence of GlcN6P. We observed a background cleavage rate from one flow cell of
0.0102 ± 0.0003 min−1 in 50 mM HEPES (pH 7.5), 150 mM KCl, and 1 mM MgCl2 (Figure 4.7)
on a PEG-passivated surface.

4.3.2

Cleavage of an ensemble of glmS ribozymes

To verify that the functional properties of the ribozyme were not compromised by surface immobilization, we measured reaction kinetics for a similar construct in solution. Measuring identicalvalues would indicate that the surface is not changing the cleavage kinetics. We excited the donor
and monitor the intensity of the acceptor fluorescence of samples of an ensemble of ribozymes in solution. In preliminary results, we obtained the cleavage rate for the RNA:DNA construct in solution
as 0.106 min−1 (Figure 4.8), which is smaller than the expected rate under similar conditions [54].
Further measurements of the cleavage rate with varying ligand and Mg2+ concentrations must be
performed to determine whether the cleavage kinetics of our glmS ribozyme construct are altered ascompared to those of a glmS ribozyme that does not contain engineered RNA:DNA handles. Also,
the ensemble measurements did not retrieve double-exponential kinetics, which could be due to alack of resolution or due to the double-exponential kinetics being a surface-dependent artifact.

111

4.4

Summary
We designed and implemented a single-molecule glmS ribozyme cleavage assay with teth-ered nanoparticles using short RNA:DNA constructs. This proof-of-concept study is a new singlemolecule measurement of glmS ribozyme cleavage. Our methods can potentially be expanded tonucleic-acid-based biosensors utilizing other aptamer domains. Previous studies have attempted tocreate variants of the glmS ribozyme that cleave upon binding of target molecules other than GlcN6P[229], raising the prospect of creating cleavable biosensors made of RNA. Use of the specificity ofriboswitches to create highly-specific biosensors is being explored [48, 49, 50].

Chapter 5

Conclusions and Future Directions

5.1

Conclusions
The studies presented here signify the first direct measurements of RecBCD·DNA destabiliza-tion dynamics. We observed large conformational destabilization dynamics of the RecBCD·DNA
complex that were on the order of multiple base pairs. This type of measurement is only possible
with an assay that can resolve subnanometer changes in position, which was accomplished using anactively-stabilized optical-trapping assay. In this thesis, we present two new, major observations
detailing RecBCD helicase mechanism. First, to conclude that the helicase was destabilizing theduplex region of the DNA substrate, it was necessary to study the conformational dynamics of
RecBCD bound to an interstrand cross-linked DNA substrate. We observed dramatically-alteredconformational dynamics in the presence of the cross link, providing strong evidence that the enzyme
actively destabilizes the DNA duplex region. Second, we measured significantly-reduced conformational dynamics in the presence of nonhydrolyzable ATP analogs. This result suggests that thedestabilization dynamics are modulated by the nucleotide-bound state of RecBCD and may be an
important aspect of helicase function, particularly in the mechanisms of active helicases.
Additionally, we developed an original single-molecule cleavage assay of the glmS ribozyme,
the first-discovered catalytic riboswitch. The work presented here is one of the first single-molecule
studies of the glmS ribozyme. This cleavage assay has the potential to be expanded to examine
other biosensors that utilize aptamer domains. Despite the fact that nucleic acids have more limited
chemical functionality than proteins, the target binding properties of aptamers can rival those113
of antibodies, and nucleic acids possess a number of advantages over antibodies, including easiersynthesis and superior transduction.

5.2

Future Directions

5.2.1

Further characterization of back-and-forth dynamics

Improved resolution of the RecBCD optical-trapping assay may be attained by repositioning
the biotin moiety. The derivative of RecBCD (RecBCD-bio) that we used in our studies has agenetically-engineered biotin moiety near the carboxy terminus of the RecD subunit. Placing thebiotin moiety at a different position on the enzyme can be achieved by posttranslational modification[230]. Modification of the enzyme would require demonstration that the new RecBCD derivativeis still functional in vivo and still exhibits helicase and DNA-dependent ATPase activities in vitro
analogous to those of wild-type RecBCD. With this approach, it may be possible to measure the mosthighly-dynamic regions of the RecBCD·DNA complex with improved spatio-temporal resolution.
Resolving the dynamics has the potential to reveal additional details of the translocation stepping
mechanism.

5.2.2

Quantification of duplex destabilization by the helicase

There are two methods of quantifying the extent to which a helicase destabilizes the junction.
In one method, the velocity of the helicase unwinding dsDNA is compared to the velocity of helicase
translocation along ssDNA. This method has been explored in several single-molecule studies [72,114, 117, 120]. In the second method, the dependence of the unwinding velocity of the helicase on
the GC-content of the double-stranded nucleic acid is examined [72, 118].
Since RecBCD will not bind to ssDNA, one strategy that could be employed in the first method
is to use DNA substrates with double-stranded regions and ssDNA gaps to induce translocation of
RecBCD on ssDNA [231]. For our RecBCD optical-trapping assay, a long DNA substrate (e.g.,
L > 500 nm) is needed to measure translocation. Such a DNA construct could be constructed with

114
double-stranded regions and a non-base-paired ssDNA region. An approach which would allow useof the second method is to design a DNA construct with tandem repeats of GC-rich and AT-richsequences. In conjunction, these methods may elucidate duplex destabilization by RecBCD.

5.2.3

Characterization of the RecB motor and RecBCD after Chi recognition

The RecBCD optical-trapping assay can be used to directly measure the stepping behaviorof the RecB motor of RecBCD. The RecBCD·DNA complex did not exhibit large conformational
dynamics when bound to DNA with a 30 -(dT)6 noncomplementary DNA tail, in spite of interaction
between the 30 DNA end with the RecB motor [191]. This begs the question of whether the destabilization dynamics are present when RecB is the lead helicase motor. Comparison of unwinding
rates of a disabled-RecD-motor mutant (RecBCDK177Q ) with unwinding rates of RecBCD before
and after Chi provided evidence that RecD (RecB) is the lead motor of RecBCD before (after) Chi
recognition [139]. In these studies, when RecD was the lead motor, RecBCD unwound DNA roughly
twice as fast as when RecB was the lead motor. Understanding the mechanistic basis of these two
superfamily-1 helicases and how they differ would bring further understanding to general helicase
mechanism.

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Appendix A

Protocols

A.1

RecBCD Tethered-Bead Assay
The following protocol was used to tether beads to a coverslip surface via a dsDNA moleculebound to a biotinylated RecBCD molecule attached to the coverslip.

RecBCD-Biotin Assay: Streptavidin-coated surface, Streptavidin-coated beads
The following buffer was used:
10 mL Wash Buffer (WB)
To a new 15 mL falcon tube add:
8.45 mL ddH2 O

final concentration:

250 µL 2 M TRIS, pH = 7.5

25 mM

10 µL 1M Mg(OAc)2

1 mM

10 µL 1M NaCl

1 mM

400 µL 10% Tween-20

0.4%

870 µL 35 mg/mL BSA (filtered)*

3 mg/mL

10 µL 1M DTT

1 mM*To make 10 mL 35 mg/mL BSA:
9.4 mL ddH2 O
200 µL 1 M Tris-Cl

132
400 µL 10% Tween-20
350 mg BSA
To filter, take two 500 µL aliquots of 35 mg/mL BSA solution and put in 0.22 um Millipore filter
tubes. Centrifuge at half-speed for 5 min. Remove the filter and take 435 µL out of each tube andput into falcon tube.
(1) Make slides and allow epoxy to dry.
(2) *Be concurrently preparing beads/DNA.*(3) Dilute 200 µL of 125 µg/mL bio-BSA solution (enough for 10 slides).
195 µL 100 mM Na-Phos pH = 7
5 µL of 5 mg/mL solution(4) Flow 20 µL per slide of 125 µg/mL Bio-BSA solution through slides.
– Sit for 20 min. in humidity chamber at room temp.
(5) Wash 2 times per slide with 200 µL WB.
– Sit for 10 min.
(6) Make 300 µL of 10 µg/mL Streptavidin (SA) (enough for 6 slides).
297 µL WB
3 µL of 1 mg/mL SA(7) Flow 50 µL per slide of 10 µg/mL SA.
– Sit for 20 min in humidity chamber at room temp.
(8) Wash 2 times per slide with 200 µL of WB.
– Sit for 10 min.

133(9) Make 40 µL of 15 nM RecBCD-bio (usually 0.45 µL of 10 µM or 34 µM RecBCD in 500µL of WB).
(10) Flow 20 µL 15 nM RecBCD-bio.
– Sit for 2 hours in humidity chamber at room temp.
(11) Wash 2 times with 200 µL of WB.
– Sit for 10 min.
(12) Flow 30 µL of beads/DNA per slide.
– Sit for 60 min in humidity chamber at room temp.
(13) Wash 2 times with 200 µL of WB.
– Sit for 10 min.
(14) View under microscope.
(15) Add 50 µL of ATP-SSB solution to slide for RecBCD to unwind the DNA.
81.4 µL WB
10 µL 100 µM ATP

10 µM ATP8.6 µL 12.8 µM SSB

1.1 µM SSB*SA beads/DNA*(1) Briefly vortex bead stock.
(2) Add 30 µL of 408 pM SA bead stock (for 440-nm diameter, 920 pM for 320-nm diameter)per 2 slides.
(3) Wash beads three times. Add 200 µL 0.4% Phos-Tween, centrifuge (5 min, 10 krpm, 400
nm bead) (7krpm, 500 nm bead, 5 min) (12 krpm, 320 nm bead, 5 min), and remove
supernatant.

134(4) Resuspend in 30 µL of WB (30 µL of 408 pM beads).
(5) Sonicate beads.
(a) Parafilm the tube before sonicating.
(b) 4 ◦ C temperature(c) 8 min pulse mode (2 s on, 2 s off)
(6) Check beads for monodispersion.
(a) Dilute 1 µL of beads in 100 µL of WB.
(b) Flow through a slide and check under microscope.
(7) Make 30 µL of 100 pM DNA per two slides.
(8) If two dilutions are necessary, 1st dilution should be in 10mM Tris-Cl (This dilution can be
kept and used for weeks.)(9) For 2nd dilution, dilute into WB. This solution can be used for ∼1 week.
(10) Mix 408 pM beads with 100 pM DNA, makes 204 pM beads and 50 pM DNA(11) Mix by pipetting.
(12) Sit for 60 min at room temperature.

135

A.2

glmS Ribozyme Tethered-Bead Assay

glmS Tethered-Bead Assay: Streptavidin-coated surface, Antidigoxigenin-coated beads
The following buffer was used:
10 mL Yeonee Buffer (YB) with Tween-20
To a new 15 mL falcon tube add:
7.625 mL DEPC-treated dH2 O (RNase-free)Final concentration:

100 µL 1 M HEPES pH 7.5

10 mM

1500 µL 1 M KCl

150 mM

400 µL 10% Tween-20

0.4%

375 µL 80 mg/mL acetylated BSA*

3 mg/mLYB and BSA solution is only good for 1 week at 4 ◦ C. Need at least 2.58 mL YB per 2 slides.
(1) Make slides and allow epoxy to dry.
(2) Be concurrently preparing beads and RNA:DNA while completing steps 3-8.
(3) Make 100 µL (per 2 slides) of 125 µg/mL Biotin BSA solution. Mix by vortexing:
97.5 µL of 100 mM Phosphate buffer (JUST Phos, no Tween!)
2.5 µL of 5 mg/mL biotinylated BSA(4) Flow 50 µL per slide of 125 µg/mL Biotin BSA solution–Let sit for 20 min in humidity chamber at room temperature.
(5) Wash slides 2x w/200 µL YB.
–Let sit for 10 min at room temperature.
(6) Make 100 µL (per 2 slides) of 10 µg/mL SA solution. Mix by vortexing:
99 µL of YB

136
1 µL of 1 mg/mL SA(7) Flow 50 µL per slide of 10 µg/mL SA solution–Let sit for 10 min in humidity chamber at room temperature.
(8) Wash slides 2x w/200 µL YB.
–Let sit for 10 min at 4 ◦ C.
*Bead-RNA:DNA preparation*(1) Prepare 40 µL 375 pM AD beads per 2 slides.
(a) Briefly vortex AD bead stock.
(b) To an epindorph tube, add enough beads to be 375 pM when diluted in 40 µL per 2
slides.
17.24 µL of 870 pM 180 nm beads per 2 slides(c) Bring up to 200 µL in Phos-Tween buffer (0.4%),(d) Centrifuge for 7 min at 14 krpm and remove supernatant.
(e) Resuspend beads in 200 µL Phos-Tween, centrifuge 7 min, remove supernatant.
(f) Resuspend in 40 µL YB.
(g) Sonicate 10 min total.
(h) Check under microscope for monodispersion.
(2) Make 40 µL 75 pM DNA per two slides.
(a) If two dilutions are necessary, 1st dilution should be in 10 mM Tris Cl (This dilution
can be kept and used for weeks.)(b) For 2nd dilution, dilute into YB. This solution can be used for ∼1 week. 3 µL of 1 nM
DNA + 37 µL YB + 2 µL SUPERase

137(3) Add DNA to bead solution.
40 µL of the 375 pM bead solution per two slides
40 µL of the 75 pM DNA solution per two slides(a) Mix by pipetting quickly 10x, then vortexing for 30 s on medium (5).
(b) Let sit 20 min at 4 ◦ C.
(4) Flow 30 µL of bead-DNA solution through each slide.
–Let sit 20 min. at 4 ◦ C.
(5) Wash slides 2x w/200 µL YB.
(6) Wash slides 2x w/200 µL No Ligand Buffer.

Appendix B

Alignment and Calibrations

B.1

Optical-Trapping Instrument
Calibrations of the precision optical-trapping instrument requires a series of steps. Most ofthese steps can be done using a polystyrene bead with diameter d = 400 nm. A free bead refers
to a bead in solution, usually an uncoated polystyrene particle in water. A stuck bead refers to abead that is stuck to a glass coverslip, typically using a buffer with high magnesium concentration
followed by a bake [87].
(1) Coarse optical alignment.
Check that the laser beams are level, not clipping on any optics, and parallel to the screw
holes in the optical table. Lenses and detectors should be aligned to the optical axis of thelaser. Fiber coupling of the laser into the fiber should be adjusted such that the output
from the AOM is imaged onto the angle-cleaved fiber tip.
(2) Optimize trap laser collimation.
Set the focal point of the trapping laser (λ = 1064 nm) to the optimal position relative tothe objective lens such that it keeps a trapped free bead at a constant axial position whenlowering the stage position. To do this, translate the 2nd lens after the fiber output in theaxial direction, and translate the output end of the fiber in the axial direction. Measure the1064 sum signal (V1064,sum ) of a trapped free bead versus stage position (zstage ), scanningfrom when the bead leaves the surface to when zstage is several microns away (Figure 2.8).

139
LabVIEW: C:\Users\Perkins\Labview \JILA\DriftofFreeBead_ASH052510.vi(subVI: C:\Users\Perkins\Labview\JILA\height JILA 2bm DAQ.vi)
IGOR:

Load general text (e.g., free223314.hg1) saved by the height program(height JILA 2bm DAQ.vi).

Examples: 052610.pxp MH #6 p.126, 061906 ARC #11 p.55
(3) Optimize the position of the bead detection laser focus.
Repeat steps as described in step 2, but using signals for the bead detection laser (λ = 785
nm) (Figure 2.8). First perform a coarse adjustment by making the image of the detection
laser focus on the CCD camera in the same image plane as the trapped bead. Measure the785 sum signal (V785,sum ) of a trapped free bead versus stage position (zstage ), scanning fromwhen the bead leaves the surface to when zstage is several microns away. Then translate the
2nd lens after the fiber output in the axial direction, and translate the output end of thefiber in the axial direction to “flatten” the bead position signal such that it stays constantas zstage increases.
(4) Position CCD camera in a plane conjugate to the trapping plane.
Determine z position at which a trapped bead leaves the coverslip surface by overlaying a
785 z scan of a free bead with that of a stuck bead (e.g., surface-to-peak ∼340 nm). Set
the CCD camera position to be at the image plane of when a stuck bead is at that position(Figure 2.7).
Examples: 052710.pxp MH #6 p.127(5) Position the focus of the fiducial mark detection laser.
Determine the trap position using the aligned 785 sum signal off a stuck bead. Measure thesum signal off the bead with the stage detection laser (λ = 850 nm). Move the 850 nm 2ndlens and fiber tip axially to optimize the 850 nm position of the focus (e.g., to be in linewith the 785 focus, or, in the case of the RecBCD assay, to be 300 nm below the trapping

140
plane).
(6) Minimize crosstalk of all lasers.
Measure the Vx , Vy , and Vz voltage signals of a stuck bead when moving the stage in x, y,
and z. Minimize the crosstalk (i.e., minimize Vx versus ystage , Vy versus xstage , Vx versus
zstage , and Vy versus zstage ). To minimize crosstalk, adjust the lateral position of the fiber
output with the 3D-translation stage upon which it is mounted.
(7) Calibrate the PZT mirrors.
Calibrate the 1064 and 850 PZT mirrors. Each PZT mirror can be rotated by orienting themirror to different screw holes in its gimbal mount.
LabVIEW: C:\Users\Perkins\Labview\JILA\Calibration Routines\CalibratePZTMirrorUsingStageAndStuckBead.vi
C:\Users\Perkins\Labview\JILA\Calibration Routines\CheckCalibratePZTMirrorUsingStageAndStuckBead.vi(8) Measure the pointing stability between the detection lasers.
Measure the differential signal between 785/850 (e.g., (V785,x − V850,x ) versus time) to measure the relative pointing stability between the 785 and 850 detection lasers. The average
standard deviation should be (0.05 nm, 0.05 nm) in (x, y) for 1-10 Hz (average 10 sec)
(Figure 2.12A).
LabVIEW: C:\Users\Perkins\Labview\Z calibration\ZCal_2_Beams_1 bead_Rezero.vi
IGOR:

Load general text (e.g., ReZero173259.dat). Run AnalyzeRezero.ipf (but do
not decimate, just smooth data) and StdDev1s.ipf.

Examples: 060410 stability.pxp MH #6 p.135, 080806 ARC #11 p.116(9) Measure stability with active stage stabilization.
Measure 785 when stabilizing with 850 for stage position lock. Use DAQ rate 10.4 kHz,

141
average 104 points (average to 100 Hz), set anti-alias filter to 2 kHz. When setting up, centerthe 785 on a stuck bead (d ∼ 400 nm) with StageDrift_1_Beam_Multi.vi and center the 850
on the same bead with BeamAlignment.vi (PZT 850, detector beam 850) after setting the
850 PZT mirror to (5 V, 5 V). In ZCal2BeamStageStream_Rezero.vi, enable stabilization(hit ReZero enabled button) and turn on position clamps in x, y, and z (click those 3
buttons). Observe 785 signals. Want 0.1-10 Hz standard deviation of (∼0.1 nm, ∼0.1 nm,∼0.1 nm) in (x, y, z). For optimal stabilization, servo the 785 and 850 both at -9 V (although

785 servo is typically -3 V).
LabVIEW: C:\Users\Perkins\Labview\Z calibration\ZCal_2_Beams_1 bead_Rezero.vi
C:\Users\Perkins\Labview\JILA\StageDrift\StageDrift_1_Beam_Multi.vi
C:\Users\Perkins\Labview\JILA\BeamAlignment.vi
IGOR:

Load general text (e.g. ReZero173259.dat). Run AnalyzeRezero.ipf and
StdDev1s.ipf.

Examples: 060410 stability.pxp MH #6 p.137, 080806 ARC #11 p.116(10) Measure pointing stability between the trap and bead detection lasers.
Measure the differential signal between the 785 and 1064 lasers, as done for the 785 and 850
lasers in step 8 (Figure 2.12B).
LabVIEW: C:\Users\Perkins\Labview\Z calibration\ZCal_2_Beams_1 bead_Rezero.vi
IGOR:

Load general text (e.g. ReZero173259.dat). Run AnalyzeRezero.ipf and
StdDev1s.ipf.

Example:060410 stability.pxp(11) Measure trap ellipticity.
Measure the position of a bead in a trap and plot the y position versus x position. Fit thedata to a line. The slope should be <0.01.
LabVIEW: C:\Users\Perkins\Labview\JILA\Yeonee\StiffnessCalbyPF&EQP.vi

142
Plot with C:\Users\Perkins\Labview\JILA\Yeonee\2DHistogram.vi
IGOR:

Load binary data.

Examples: 040310 ellipticity.pxp MH #6 p.138(12) Calibrate trap stiffness versus servo voltage.
Use 400 nm uncoated bead to measure trap stiffness using the power spectrum and equipartition methods. They should agree to within 10%. Check the calibration with 3 beads. The
error should be within 10%. Typical r200stiff-to-volt values were (96, -.04, 49, -.04).
LabVIEW: C:\Users\Amanda\Labview\Calibrations\Stiffness Calibrations\Pow vs Stiff_PSandEQP_ PositonLockOpt_AECv5_mh.vi(check for Servo 1-9 V)
C:\Users\Perkins\Labview\JILA\Calibration Routines\StiffnessCalCheck.vi
IGOR:

Run plot_ktrap.ipf (loads and plots data from .kvV file or .pve file) andAnalyzeStiffCheck.ipf.

(13) Measure trap stiffness versus height.
The trap stiffness should only decrease by 15% 1 µm above the surface.
LabVIEW: C:\Users\Perkins\Labview\JILA\Yeonee\StiffnessCalbyPF&EQP.vi (checks
one voltage)
Example:040506 ARC #11 p.9, 060510 MH #6 p.140(14) Check 785 sensitivity versus z of trapped bead.

(15) Calibrate trap stiffness versus voltage using PS, EQP, and drag.
Vary voltage from 1 V to 9 V.
(16) Check linearity of trap.
Use drag to measure the trap linearity of the force versus displacement of a free bead. This

143
program moves the stage at varying velocity to drag a free bead out of the trap. Measure
the slope of the sample (dz/dx and dz/dy). Settings: 800 µm/s, number of increments 30,∼100 ms time.

LabVIEW: C:\Users\Perkins\Labview\calibrate_free_bead.vi(17) Check stiffness for 320 SA beads.
Should get r160stiff-to-volt values of (110, 0, 46, 0).
(18) Check DNA tethers for good L0 , Lp , Foffset .
Use 2 µm DNA, 400 nm beads (AD coated) with bio-BSA/SA surface.
LabVIEW: C:\Users\Perkins\Labview\JILA\DNA stretching\DNA_stretching.llb\DNA_stretching.vi
IGOR:

Procedure_GetExtensionandForce.ipf, ModifiedMarkoSiggitFit.ipf

Example:060710.pxp(19) Optimize 1064 fiber coupling.
See ARC thesis (Appendix E.11 “How to guide: fiber alignment”). Optimize at V1064,AOM =0.6 V. Get >7.25 V on PD.
(20) Check 1064 Power versus servo voltage.
Measure the power of the 1064 laser for 1064 servo voltage 1-9 V and fit to a line. Should
get 76 mW/Volt. Use large power head meter set right before beam enters microscope, butmake sure sensor is not at the focus of the telescope lens.
Example:061210.pxp(21) Measure free bead versus time.

144
LabVIEW: C:\Users\Perkins\Labview\JILA\freebead meas\Measure freebead with steps.vi
IGOR:

AnalyzeFreeBead.ipf(22) Measure stability of a DNA tether, L versus time.

LabVIEW: C:\Users\Perkins\Labview\JILA\RecBCD\RecBCDPulling_v1.llb\RecBCDPulling_v1.vi

Appendix C

Data

The data of individual molecules often contain some level of heterogeneity. For that reason,
the individual records of RecBCD bound to DNA are included here.

146

Figure C.1: DNA tethers. (A) DNA contour length versus time with corresponding histograms.
Data were taken at F = 6 pN and smoothed with a Savitzky-Golay window to 10 Hz (grey) and 1
Hz (black ). (B) Integrated fluctuations of the records in A.

147

Figure C.2: 50 -(dT)10 -tailed DNA tethers. (A) DNA contour length versus time with corresponding
histograms. Data were taken at F = 6 pN and smoothed with a Savitzky-Golay window to 10 Hz(yellow ) and 1 Hz (olive). (B) Integrated fluctuations of the records in A.

148

Figure C.3: Control data: RecBCD bound to blunt-ended DNA. (A) DNA contour length versus
time with corresponding histograms. Data were taken at F = 6 pN and smoothed with a SavitzkyGolay window to 10 Hz (light blue) and 1 Hz (blue). (B) Integrated fluctuations of the records in
A.

149

Figure C.4: Control data: RecBCD bound to partially unwound DNA in the absence of ATP. (A)DNA contour length versus time with corresponding histograms. Data were taken at F = 6 pN andsmoothed with a Savitzky-Golay window to 10 Hz (pink ) and 1 Hz (red ). (B) Integrated fluctuationsof the records in A.

150

Figure C.5: RecBCD bound to 50 -(dT)6 -tailed DNA. (A) DNA contour length versus time withcorresponding histograms. Data were taken at F = 6 pN and smoothed with a Savitzky-Golay
window to 10 Hz (light yellow ) and 1 Hz (dark yellow ). (B) Integrated fluctuations of the records
in A.

151

Figure C.6: RecBCD bound to 50 -(dT)8 -tailed DNA. (A) DNA contour length versus time withcorresponding histograms. Data were taken at F = 8 pN and smoothed with a Savitzky-Golay
window to 10 Hz (light purple) and 1 Hz (deep purple). (B) Integrated fluctuations of the records
in A.

152

Figure C.7: RecBCD bound to 50 -(dT)10 -tailed DNA. (A) DNA contour length versus time withcorresponding histograms. Data were taken at F = 6 pN and smoothed with a Savitzky-Golay
window to 10 Hz (light brown) and 1 Hz (brown). (B) Integrated fluctuations of the records in A.

153

Figure C.8: RecBCD bound to 50 -(dT)20 -tailed DNA. (A) DNA contour length versus time withcorresponding histograms. Data were taken at F = 20 pN and smoothed with a Savitzky-Golay
window to 10 Hz (light green) and 1 Hz (green). (B) Integrated fluctuations of the records in A.

154

Figure C.9: RecBCD bound to 50 -(dT)10 -tailed DNA in the presence of ADP·BeFx . (A) DNA
contour length versus time with corresponding histograms. Data were taken at F = 6 pN andsmoothed with a Savitzky-Golay window to 10 Hz (light purple) and 1 Hz (purple). (B) Integrated
fluctuations of the records in A.

155

Figure C.10: RecBCD bound to 50 -(dT)10 -tailed DNA in the presence of AMPPNP. (A) DNA
contour length versus time with corresponding histograms. Data were taken at F = 6 pN andsmoothed with a Savitzky-Golay window to 10 Hz (light yellow ) and 1 Hz (yellow ). (B) Integrated
fluctuations of the records in A.

156

Figure C.11: RecBCD bound to 50 -(dT)10 -tailed DNA in the presence of ADP + Pi . (A) DNA
contour length versus time with corresponding histograms. Data were taken at F = 6 pN andsmoothed with a Savitzky-Golay window to 10 Hz (light gray) and 1 Hz (gray). (B) Integrated
fluctuations of the records in A.

157

Figure C.12: RecBCD bound to 50 -(dT)10 -tailed DNA in the presence of ADP. (A) DNA contour
length versus time with corresponding histograms. Data were taken at F = 6 pN and smoothedwith a Savitzky-Golay window to 10 Hz (light blue) and 1 Hz (sky blue). (B) Integrated fluctuationsof the records in A.

158

Figure C.13: RecBCD bound to 50 -(dT)10 -tailed DNA in the presence of ATPγS. (A) DNA contour
length versus time with corresponding histograms. Data were taken at F = 6 pN and smoothedwith a Savitzky-Golay window to 10 Hz (sea green) and 1 Hz (teal ). (B) Integrated fluctuations of
the records in A.

159

Figure C.14: RecBCD bound to AT-rich 50 -(dT)10 -tailed DNA at 2 pN. (A) DNA contour length
versus time with corresponding histograms. Data were taken at F = 2 pN and smoothed with a
Savitzky-Golay window to 10 Hz (light blue) and 1 Hz (cyan). (B) Integrated fluctuations of therecords in A.

160

Figure C.15: RecBCD bound to AT-rich 50 -(dT)10 -tailed DNA at 4 pN. (A) DNA contour length
versus time with corresponding histograms. Data were taken at F = 4 pN and smoothed with a
Savitzky-Golay window to 10 Hz (light blue) and 1 Hz (light turquoise). (B) Integrated fluctuationsof the records in A.

161

Figure C.16: RecBCD bound to AT-rich 50 -(dT)10 -tailed DNA at 6 pN. (A) DNA contour length
versus time with corresponding histograms. Data were taken at F = 6 pN and smoothed with a
Savitzky-Golay window to 10 Hz (light blue) and 1 Hz (turquoise). (B) Integrated fluctuations of
the records in A.

162

Figure C.17: RecBCD bound to AT-rich 50 -(dT)10 -tailed DNA at 10 pN. (A) DNA contour length
versus time with corresponding histograms. Data were taken at F = 10 pN and smoothed with a
Savitzky-Golay window to 10 Hz (light blue) and 1 Hz (dark turquoise). (B) Integrated fluctuationsof the records in A.

163

Figure C.18: RecBCD bound to mixed-sequence 50 -(dT)10 -tailed DNA at 2 pN. (A) DNA contour
length versus time with corresponding histograms. Data were taken at F = 2 pN and smoothedwith a Savitzky-Golay window to 10 Hz (light yellow-orange) and 1 Hz (dark yellow-orange). (B)Integrated fluctuations of the records in A.

164

Figure C.19: RecBCD bound to mixed-sequence 50 -(dT)10 -tailed DNA at 4 pN. (A) DNA contour
length versus time with corresponding histograms. Data were taken at F = 4 pN and smoothedwith a Savitzky-Golay window to 10 Hz (light orange) and 1 Hz (dark orange). (B) Integrated
fluctuations of the records in A.

165

Figure C.20: RecBCD bound to mixed-sequence 50 -(dT)10 -tailed DNA at 10 pN. (A) DNA contour
length versus time with corresponding histograms. Data were taken at F = 10 pN and smoothedwith a Savitzky-Golay window to 10 Hz (light brown) and 1 Hz (dark brown). (B) Integrated
fluctuations of the records in A.

166

Figure C.21: RecBCD bound to GC-rich 50 -(dT)10 -tailed DNA at 2 pN. (A) DNA contour length
versus time with corresponding histograms. Data were taken at F = 2 pN and smoothed with a
Savitzky-Golay window to 10 Hz (light rose) and 1 Hz (rose). (B) Integrated fluctuations of therecords in A.

167

Figure C.22: RecBCD bound to GC-rich 50 -(dT)10 -tailed DNA at 4 pN. (A) DNA contour length
versus time with corresponding histograms. Data were taken at F = 4 pN and smoothed with a
Savitzky-Golay window to 10 Hz (light pink ) and 1 Hz (pink ). (B) Integrated fluctuations of therecords in A.

168

Figure C.23: RecBCD bound to GC-rich 50 -(dT)10 -tailed DNA at 6 pN. (A) DNA contour length
versus time with corresponding histograms. Data were taken at F = 6 pN and smoothed with a
Savitzky-Golay window to 10 Hz (light pink ) and 1 Hz (dark pink ). (B) Integrated fluctuations of
the records in A.

169

Figure C.24: RecBCD bound to GC-rich 50 -(dT)10 -tailed DNA at 10 pN. (A) DNA contour length
versus time with corresponding histograms. Data were taken at F = 10 pN and smoothed with a
Savitzky-Golay window to 10 Hz (light magenta) and 1 Hz (magenta). (B) Integrated fluctuationsof the records in A.

170

Figure C.25: RecBCD bound to AT cross-linked 50 -(dT)10 -tailed DNA. (A) DNA contour length
versus time with corresponding histograms. Data were taken at F = 6 pN and smoothed with a
Savitzky-Golay window to 10 Hz (lighter blue) and 1 Hz (light blue). (B) Integrated fluctuations of
the records in A.

171

Figure C.26: RecBCD bound to GC cross-linked 50 -(dT)10 -tailed DNA. (A) DNA contour length
versus time with corresponding histograms. Data were taken at F = 6 pN and smoothed with a
Savitzky-Golay window to 10 Hz (light green) and 1 Hz (green). (B) Integrated fluctuations of therecords in A.

172

Figure C.27: RecBCD bound to Low-AT 50 -(dT)10 -tailed DNA. (A) DNA contour length versus
time with corresponding histograms. Data were taken at F = 6 pN and smoothed with a SavitzkyGolay window to 10 Hz (light orange) and 1 Hz (orange). (B) Integrated fluctuations of the records
in A.

173

Figure C.28: RecBCD bound to Low-GC 50 -(dT)10 -tailed DNA. (A) DNA contour length versus
time with corresponding histograms. Data were taken at F = 6 pN and smoothed with a SavitzkyGolay window to 10 Hz (light green) and 1 Hz (green). (B) Integrated fluctuations of the records
in A.