Single-Molecule Kinetic and Thermodynamic Studies of Cosolute-Influenced Nucleic Acid
Conformational Transitions
by
Erik Dylan Holmstrom
B.A., Willamette University, 2008

A thesis submitted to the
Faculty of the Graduate School of the
University of Colorado in partial fulfillmentof the requirement for the degree ofDoctor of Philosophy
Biochemistry2014This thesis entitled:
Single-Molecule Kinetic and Thermodynamic Studies of Cosolute-Influenced Nucleic Acid
Conformational Transitions
written by Erik Dylan Holmstrom
has been approved for the Department of Chemistry and Biochemistry

David J. Nesbitt

Robert T. Batey
Date
The final copy of this thesis has been examined by the signatories, and weFind that both the content and the form meet acceptable presentation standards
Of scholarly work in the above mentioned discipline.

Holmstrom, Erik Dylan (Ph.D., Biochemistry)Single-Molecule Kinetic and Thermodynamic Studies of Cosolute-Influenced Nucleic Acid
Conformational Transitions
Thesis directed by Professor David J. Nesbitt

Over the last 40 years the number of biochemical functionalities attributed to nucleic acids
has increased tremendously. This diverse array of chemical functionality is intimately coupled tothe spatial arrangement of atoms associated with these molecules. The three-dimensional
structures and functions of nucleic acids are known to be dependent on the concentration andidentity of solutes in solution. These nucleic acid cosolutes can be as simple and universal asatomic metal cations that favorably interact with the negatively charged phosphate backbone of
nucleic acids and resulting in stabilization of electronegatively dense conformations.
Alternatively, they may be complex organic molecules that are able to promote conformationaltransitions in certain RNA sequences responsible for regulating gene expression. Understanding
the biophysical principles responsible for these cosolutes-influenced conformational transitions
represents the primary objective of this work.
Single-molecule fluorescence resonance energy transfer (smFRET) is a technique that iswell suited for this goal because it provides the ability to watch individual nucleic acids undergocosolute-influenced conformational transitions in real-time. These experimental results provide
direct access to rate and equilibrium constants for any nucleic acid of interest. The temperaturedependence of these smFRET experiments can be used to partition the free energy change
associated with these conformational transitions into the enthalpic and entropic components. The
kinetic and thermodynamic information acquired from these experiments, in conjunction withiii

existing structural models, is used to describe the folding pathways and free energy landscapes
associated with these nucleic acids.
The types of nucleic acid conformational transitions described in this thesis are as follows:
the GAAA Tetraloop Receptor interaction, a 21 bp DNA duplex, and 8bp DNA duplex, aminimal version of the human Telomerase RNA pseudoknot, and a vitamin-B12 riboswitch. The
cosolutes investigated for these nucleic acid constructs include: MgCl2, KCl, NaCl, urea,
trimethylamine oxide, and vitamin-B12. Lastly, this thesis documents the development of anadvanced heating technique used to conduct many of the temperature-dependent experiments
associated with the aforementioned nucleic acids.

iv

Acknowledgments
This thesis would not exist without never-ending support, encouragement, and inspirationfrom a number of individuals. The following is a brief, and almost certainly incomplete, list ofpeople that deserve some acknowledgment for their role in getting me here to write this thesis.
First and foremost, I would like to thank my immediate family, especially my parents—
Roy and Sue Holmstrom. Throughout my entire life they have remained positive and enthusiastic
about all of my scientific interests. It is their parental guidance and financial assistance that has
provided me the opportunity to attend graduate school. Lindsay Holmstrom has been a wonderfulsister, and deserves credit for being patient with my long-winded answers to her chemical andbiological questions. As a nurse, she has also continually fostered my interest in issues related tohuman health.
It was in elementary school where I remember my first prolonged exposure to the naturaland physical sciences and for that I would like to thank my third grade teacher—Mrs. Walley.
Early exposure to the sciences certainly played an important role in setting me on the path that
led to this thesis. I am grateful for Mr. Micheletto, my seventh grade science teacher, who wasalways able to make science interesting and applicable to the real world. However, it was not
until high school that I fully realized my devotion to the sciences in the classes of: Mr. Honzel(earth science), Mr. McMaster (anatomy and physiology), Mrs. Prather (biology), Ms. Smith(chemistry), and Mr. Govertson (physics). I must also acknowledge Mr. Little (calculus) andMrs. Umba (statistics) for demonstrating how fundamental math is in describing the world on
which we live.
This amazing collection of pre-collegiate experiences primed me for an enlightening
tenure as an undergraduate student. I would like to extend my gratitude to the many mentors and!

v

advisors I had during those four years: (i) Myles Jackson—history and philosophy of science(Willamette University), (ii) Susan Kephart— biology (Willamette University), (iii) Sarah
Kirk—chemistry (Willamette University), (iv) Kent Sugden and his wife Brooke Martin—summer research (University of Montana) and (v) David Johnson—summer internship(GlaxoSmithKline).
Myles is certainly the most memorable educator I have ever interacted with and only
those who have had the opportunity to experience his wisdom would know why. Susan andSarah continually encouraged me to actually apply my talents to my studies and encouraged me
to consider the possibility of graduate school. Kent Sugden, Brooke Martin, and David Johnson
gratefully opened up their laboratories, which allowed me to experience real scientific research.
It was Kent who first piqued my interest in nucleic acids and, at the last minute, encouraged me
to apply to the University of Colorado at Boulder for graduate school.
My first exposure to the fundamental ideas behind the work contained within this thesis
was through my eventual graduate student mentor, Julie Fiore, at a poster session during thegraduate student recruitment weekend in the spring of 2008. If I could only acknowledge one
person for his or her contributions to this thesis it would be hard to say thanks to anyone otherthan Julie. She taught me exactly what I needed to know to prosper as a graduate student andprovided a great source of scientific motivation and information. Thank you Julie! Tom Baker
also deserves special mention for his omnicious presence in graduate school. It didn’t take longfor me to understand why everyone in the department came to Tom for answers to theirquestions. The lab was never the same after he graduated. Both Tom and Julie were first classsenior graduate students during my first few years in Boulder.

!

vi

I would also like to thank all of the other lab mates I have had over the years for theirmemorable contributions to my graduate school experience: Nick Dupuis for Friday afternoon
experiments, Kevin Early for listening to KEXP, Grant Buckingham for pushup contests, andAndy Gisler for being the lab general, Mike Ziemkiewicz for lunchtime volleyball, Mario
Vieweger for the Top 40, Larry Fiegland for help with Lab Windows, Andrej Grubisic for beinga great neighbor, Rob Roscioli for darts at the Sundown Saloon, Danny Bell for remaining
incredibly calm during some contentious group presentations, Mia Zutz for the baked goods,
Lora Nugent-Glandorf for showing me the TIRF microscope, Monika Gruetter for continually
asking for company on coffee breaks (even though I don’t drink coffee), Dan Nelson for wearing
clogs in the lab, Mel Roberts for hockey games, Erin Sharp-Willams for the birthday cakes,
Volker Schweikhard for the computer and workstation, Chih-Hsuan Chang for learning how to
ice skate with the lab, and Fang Wang for continually talking about organizing group activities.
I would like to acknowledge the 2008 class of biochemistry graduate students, most ofwhom I consider friends rather than classmates. I would like to thank the many colleagues I have
had the opportunity to work with over the years: Rob Batey, Andrew Garst, Jake Polaski, Jeremy
Trausch, Joe Falke, Kene Piasta, Yun-xing Wang, and Yu Li. It is also worth noting a few of thepeople who have provided useful feedback during the many struggles and obstacles of graduate
school: Laura Johnson, Dan Hickstein, Dan Adams, Kevin Dean, Allison Churnside, Thayne
Dickey, Art Pardi, Jeff Swan, Mike Yarus, Tom Cech, Cara Florence, and Debbie Wutke.
My graduate school experience has been significantly enriched by the NIH/ University ofColorado Molecular Biophysics program, which is passionately run by Joe Falke with additional
support provided by Lin Pharris. JILA—the jointly run set of research labs between the National
Institutes of Standards and Technology (NIST) and the University of Colorado at Boulder—has!

vii

been the home of my graduate research for the past 6 years. This building is not only populated
by a number of Noble laureates and MacArthur fellows, but also by some of the best staff, andadministration on campus. Specific mention is essential for Julie Philips and Brad Baxley from
SRO, and J.R. Wraith, Philip Kao, Jim Mckown, Alan Dunwell, and Peter Ruprecht from
Computing. The JILA electronics and instruments shops also need acknowledgement, notably:
James Fung-a-Fat, Carl Sauer, Hans Green, Todd Asincar, and Kim Hagen. I would like to thankJeffrey Sauter, Brian Lynch and Randall Holliness from the purchasing office for handing theunusually complex process associated with purchasing nucleic acids as well as a couple ofhelpful people from JILA administration—Maryly Dole and Agnieszka Lynch.
Thanks to my oral examination committee—Rob Batey, Art Pardi, and Amy Palmer—for
ensuring that I had the necessary skills to advance to PhD candidacy. Thanks to my thesis
Committee for their assistance and support while I was preparing to graduate: Norm Pace, Dick
McIntosh, Joe Falke, Rob Batey, and, most importantly, my research Advisor—David Nesbitt.
David shares a diverse collection of qualities with many of the most influential people in
my life. Like my parents did when I was a child, David has provided me with financial support.
Like my seventh grade science teacher, David has shown me the importance of science in thereal world. Like my high school math teachers, David has stressed the importance of math indescribing scientific observations. Like my undergraduate chemistry and biology advisors, David
has continually asked me to “raise the bar” and apply my talents to my research. As my graduate
research advisor, David taught be to be a strong critic of my own work. His vast reservoir ofscientific wisdom and intuition served as a great Litmus test for how well I understood my own
research. Most importantly, he provided me with a graduate school experience unlike any otherin the world and for that he deserves special acknowledgment.

!

viii

Last, but certainly not least, I would like to thank the single most important person in mylife—my partner and best friend—Jennifer McWilliams. She has been by my side for my entire
graduate career, which inevitably involves listening to me talk about my research to myself late
at night while staring at the computer. Her undying support and encouragement has allowed me
to preserver though the trials and tribulations of graduate school. Jen’s warm compassionate
heart and cheerful sense-of-humor have always been able to brings a smile to my face, no matter
what was holding me down. Most importantly, she has provided a source of balance in my life
like none other and for that I am forever grateful.

!

ixTable of Contents
Chapter 1:
Introduction .................................................................................................................................. 1
1.1
1.2
1.3
1.4
1.5

Nucleic Acids ............................................................................................................. 1
Nucleic Acid Conformational Transitions ................................................................. 5
Cosolute-Influenced Nucleic Acid Conformational Transitions ............................... 7
Single-Molecule Fluorescence ................................................................................. 11
References ................................................................................................................ 16

Chapter 2:
Experimental Methods .............................................................................................................. 19
2.1 Experimental Observation ....................................................................................... 19
2.1.1 Molecular Fluorescence ............................................................................ 19
2.1.2 Forster Resonance Energy Transfer (FRET) ............................................ 22
2.2 Single-Molecule Confocal Microscope ................................................................... 25
2.2.1 Excitation Path ......................................................................................... 25
2.2.2 Microscope and Sample Stage ................................................................. 27
2.2.3 Emission path ........................................................................................... 29
2.2.4 TCSPC Detection ...................................................................................... 30
2.2.5 Temperature Control ................................................................................. 30
2.2.5.1 Bulk Heating Methods ............................................................ 31
2.2.5.2 Infrared-based Heating............................................................ 33
2.3 Sample Preparation .................................................................................................. 34
2.3.1 Sample Holder .......................................................................................... 34
2.3.2 Sample of Interest ..................................................................................... 36
2.4 Data Collection and Analysis................................................................................... 38
2.4.1 Single-Molecule Freely Diffusing ............................................................ 38
2.4.2 Single-Molecule Surface Immobilized ..................................................... 41
2.5 Single-Molecule Construct Design .......................................................................... 43
2.6 References ................................................................................................................ 45
Chapter 3:
Real-Time IR Overtone Laser Control of Temperature in Picoliter H2O Samples: “NanoBathtubs” for Single Molecule Microscopy .............................................................................. 47
3.1
3.2
3.3
3.4
3.5

Introduction .............................................................................................................. 47
Results and Discussion ............................................................................................ 49
Conclusions .............................................................................................................. 56
Experimental Methods ............................................................................................. 57
References ................................................................................................................ 58

x

Chapter 4:
Pulsed Infrared-Heating Studies of Single Molecule DNA Duplex Dissociation Kinetics andThermodynamics........................................................................................................................ 60
4.1 Introduction .............................................................................................................. 60
4.2 Overtone Infrared Laser Based Heating Technique................................................. 64
4.2.1 In Situ Measurement and Dynamic Range of Accessible Temperatures .. 65
4.2.2 High Resolution Spatial Control of Temperature ..................................... 67
4.2.3 Rapid Temperature Modulation ................................................................ 69
4.3 A Test Application: Single Molecule Kinetics of DNA Duplex Dissociation ........ 73
4.3.1 Laser Heating Induced Duplex DNA Dissociation Kinetics .................... 74
4.4 Duplex Dissociation Model ..................................................................................... 81
4.5 Conclusions .............................................................................................................. 85
4.6 Experimental Methods ............................................................................................. 86
4.6.1 Fluorescent Oligonucleotides.................................................................... 86
4.6.2 Sample Preparation ................................................................................... 87
4.6.3 Fluorescence Microscope.......................................................................... 87
4.6.4 Duplex Dissociation Experiments............................................................. 87
4.7 References ................................................................................................................ 89
Chapter 5:
Thermodynamic Origins of Monovalent-Facilitated RNA Folding ...................................... 94
5.1 Introduction .............................................................................................................. 94
5.2 Molecular Model ...................................................................................................... 96
5.3 Results ...................................................................................................................... 99
5.3.1 Folding Kinetics and Standard State Free Energy of Folding .................. 99
5.3.2 Standard State Enthalpy and Entropy of Folding ................................... 101
5.3.3 Transition State Thermodynamics .......................................................... 105
5.3.4 Temperature Dependence of kfold ............................................................ 107
5.4 Discussion .............................................................................................................. 112
5.4.1 Folding Pathways and Ion Atmospheres................................................. 112
5.4.2 Folding in a Free Energy Neutral Monovalent Ion Atmosphere ............ 113
5.4.3 Folding in a “Free Energy Favorable” Monovalent Ion Atmosphere ..... 115
5.4.4 Dependence on Monovalent Cation Identity .......................................... 120
5.5 Conclusion ............................................................................................................. 121
5.6 Experimental Methods ........................................................................................... 124
5.6.1 Experimental Apparatus.......................................................................... 124
5.6.2 Temperature Control ............................................................................... 124
5.6.3 RNA and Sample Preparation ................................................................. 125
5.7 Acknowledgment ................................................................................................... 125
5.8 References .............................................................................................................. 126

xiChapter 6:
Kinetic and Thermodynamic Origins of Osmolyte-Influenced Nucleic Acid Folding ...... 131
6.1 Introduction ............................................................................................................ 131
6.2 Results .................................................................................................................... 134
6.2.1 Folding in the Absence of Osmolytes ..................................................... 136
6.2.2 Osmolyte-Dependent Folding Kinetics................................................... 137
6.2.3 Osmolyte-Dependent Folding Free Energies (m-value analysis) ........... 139
6.2.4 Osmolyte-Dependent Thermodynamics for the 8 bp DNA Duplex ....... 140
6.2.5 Osmolyte-Dependent Thermodynamics for the TL-TLR ....................... 143
6.3 Discussion .............................................................................................................. 144
6.3.1 TMAO-Dependent Folding Free Energies.............................................. 145
6.3.2 Urea-Dependent Folding Free Energies.................................................. 146
6.3.3 Kinetic Origins of Osmolyte-Influenced Nucleic Acid Folding ............. 146
6.3.4 Osmolyte-Dependent Entropy Changes for the 8bp DNA Duplex ......... 148
6.3.5 Osmolyte-Dependent Entropy Changes for the TL-TLR ....................... 149
6.4 Conclusions ............................................................................................................ 151
6.5 Experimental Methods ........................................................................................... 153
6.5.1 Nucleic Acids .......................................................................................... 153
6.5.2 Solution and Sample Preparation ............................................................ 154
6.5.3 smFRET Microscope .............................................................................. 155
6.6 References .............................................................................................................. 157
Chapter 7:
Single Molecule FRET Studies of the Human Telomerase RNA Pseudoknot:
Temperature/Urea Dependent Folding Kinetics and Thermodynamics ............................ 161
7.1 Introduction ............................................................................................................ 161
7.2 Results .................................................................................................................... 164
7.2.1 Freely Diffusing Burst Measurements .................................................... 164
7.2.2 Surface Immobilized Images .................................................................. 168
7.2.3 Surface Immobilized Time Trajectories ................................................. 170
7.2.4 van’t Hoff Thermodynamics ................................................................... 174
7.3 Discussion .............................................................................................................. 177
7.3.1 Folding Free Energies ............................................................................. 177
7.3.2 Urea-dependent Free Energies and Rate Constants ................................ 180
7.3.3 Urea-Dependent Entropies and Enthalpies ............................................. 181
7.4 Conclusions ............................................................................................................ 183
7.5 Experimental Methods ........................................................................................... 184
7.5.1 Construct Design ..................................................................................... 184
7.5.2 Single-molecule Microscope .................................................................. 186
7.5.3 Freely Diffusing smFRET Experiments ................................................. 187
7.5.4 Surface Immobilized smFRET Experiments ........................................... 189
7.5.5 smFRET Temperature Control ............................................................... 190
7.5.6 smFRET Buffer Conditions .................................................................... 190

xii

7.6 References .............................................................................................................. 191
Chapter 8:
Vitamin B12-facilitates formation of a regulatory RNA kissing loop ................................. 196
8.1 Introduction ............................................................................................................ 196
8.2 Results .................................................................................................................... 200
8.2.1 KL docking equilibrium in the absence of ligand ................................... 201
8.2.2 KL docking equilibrium in the presence of ligand ................................. 204
8.2.3 Ligand binding properties ....................................................................... 205
8.2.4 CBL-dependence of the KL docking equilibrium .................................. 209
8.2.5 KL docking kinetics in the absence of ligand ......................................... 211
8.2.6 KL docking kinetics in the presence of ligand........................................ 213
8.2.7 Cell-based CBL-dependent expression of GFP ...................................... 216
8.3 Discussion .............................................................................................................. 218
8.3.1 Kissing Loop docking dynamics ............................................................. 218
8.3.2 KL docking in the presence of a macromolecular stabilizing agent ....... 219
8.3.3 Ligand Recognition ................................................................................. 221
8.3.4 Mechanism of regulation for the env8 hydroxocobalamin riboswitch .. 222
8.4 Conclusions ........................................................................................................... 223
8.5 Experimental Methods .......................................................................................... 224
8.5.1 Fluorescent Construct Design ................................................................ 224
8.5.2 Freely-Diffusing smFRET ..................................................................... 226
8.5.3 Ensemble Fluorometry ........................................................................... 226
8.5.4 Surface-Immobilized smFRET .............................................................. 227
8.5.5 in vivo Regulation of GFP Expression ................................................... 227
8.6 References .............................................................................................................. 229
Bibliography ............................................................................................................................. 233

xiii

List of Tables
Table 2.1 ....................................................................................................................................... 22
Table of approximate photophysical properties associated with four fluorophorescommonly used throughout this thesis. All reported values are for free dye moleculesin buffered aqueous solutions, expect for Cy3-DNA*, where the conjugation to anucleic acid significantly increases the quantum yield and fluorescence lifetime of thefluorophore.
Table 6.1 ..................................................................................................................................... 142
Values for the van’t Hoff analysis of the osmolyte-influenced folding nucleic acids.
Additional information about the methods of data analysis for the van’t Hoff plots canbe found in Section 6.3.4. See Figure 6.6 for experimental data.
Table 7.1 ..................................................................................................................................... 173
Rate and equilibrium constants for the a) wild-type and b) mutant forms of the humantelomerase RNA pseudoknot under varying urea concentrations. For both RNA
pseudoknot constructs, the rate constant for forming the pseudoknot from the hairpin
conformation is largely insensitive to urea concentration.
Table 7.2 ..................................................................................................................................... 176
Thermodynamics parameters from the single-molecule van’t Hoff analyses. a) Entropy
and enthalpy changes for the wild-type human telomerase pseudoknot at various urea
concentrations. ΔH° and ΔS° are (i) insensitive or (ii) linearly dependent on urea
concentration, respectively. Together they predict folding thermodynamics under nondenaturing conditions. b) Enthalpy and entropy changes for the dyskeratosis congenita
mutant form of the human telomerase RNA pseudoknot under non-denaturingconditions.

xiv

List of Figures
Chapter 1:
Figure 1.1 ...................................................................................................................................... 2
Structural components of a nucleic acid. (a) Hydrogen bonding pattern between pairsof guanine and cytosine nucleobases. (b) Hydrogen bonding pattern between pairs ofadenine and thymine nucleobases in DNA. (c) Hydrogen bonding pattern between
pairs of adenine and uracil nucleobases in RNA. (d) Schematic representation of thephosphate, sugar, and base constituents in the context of duplex deoxyribonucleic acid.
Figure 1.2 ...................................................................................................................................... 3
Central Dogma of Molecular Biology. Schematic representation of the linear transfer
of genetic information in a cell. Pointed and blunted arrows represent some of themany ways that RNA can influence the processing of this genetic information.
Figure 1.3 ...................................................................................................................................... 5
Hierarchical folding of RNA. Example depicts the major structural transitions
associated with the formation of a tRNA molecule. Colors correspond to base paired
regions within the oligonucleotide.
Figure 1.4 ...................................................................................................................................... 6
Cartoon graphics depicting the numerous oligonucleotide conformation transitions
discussed in the chapters of this thesis.
Figure 1.5 ...................................................................................................................................... 9
Chemical structures of two osmolytes known to significantly influence theconformational transitions of nucleic acids.
Figure 1.6 .................................................................................................................................... 13
Fluorescence Resonance Energy Transfer (FRET). An excited donor fluorophore(green) can transfer energy to a nearby acceptor fluorophore (red) in a distancedependent process. When the donor and acceptor are proximal, energy transfer isefficient; when they are distal, energy transfer is poor.
Chapter 2:
Figure 2.1 .................................................................................................................................... 21
Jablonski energy level diagram depicting absorption (excitation) and emission(fluorescence) of photons.
(http://www.microscopyu.com/articles/fluorescence/fret/fretintro.html)
Figure 2.2 .................................................................................................................................... 24
Jablonski energy level diagram depicting Fluorescence Resonance Energy Transfer(FRET). (http://www.microscopyu.com/articles/fluorescence/fret/fretintro.html)
Figure 2.3 .................................................................................................................................... 25
Distance dependence of FRET efficiency (EFRET) for a fictitious biological molecule.
Separation of the donor and acceptor fluorophores results in less efficient energy

xvtransfer.
Figure 2.4 .................................................................................................................................... 26
Cartoon cut-away diagram of the single-molecule microscope system used to conduct
the research associated with this thesis.
Figure 2.5 .................................................................................................................................... 28(left and middle) Visual depictions of the Airy disc associated with a diffractionlimited focus. (right) Graphical representation of the normalized radial intensity
distribution (red solid line, I(x)/Io) and the relative encircled power (blue dotted line,
P(x)/Po). Note x = kasin(θ) =2πrN.A./λ, as used in Eq. 2.7.
(http://en.wikipedia.org/wiki/Airy_disk).
Figure 2.6 .................................................................................................................................... 28
Sample surface image of immobilized fluorescence molecules using the 3-D nanopositioning peizo-electric scanning stage (not shown).
Figure 2.7 .................................................................................................................................... 31
Diagram of the original, bulk, heating capabilities established by Dr. Julie Fiore (stage
not shown).
Figure 2.8 .................................................................................................................................... 33
Cut-away cartoon of the infrared-based heating apparatus coupled to the existing
inverted confocal fluorescence microscope.
Figure 2.9 .................................................................................................................................... 35
Image of the three types of sample holders used for fluorescence studies. See Section
2.3.1 for details.
Figure 2.10 .................................................................................................................................. 38
Schematic representation (not-to-scale) of the commonly used biotin- streptavidin
surface immobilization technique. Approximately one out of ten BSA(blue ovals)
molecules contains a biotin (black triangle) moiety. Streptavidin (purple rectangle) has
four binding sites for biotin, which can be used to immobilize fluorescent nucleic acids
to the glass coverslip.
Figure 2.11 .................................................................................................................................. 39
Schematic for freely-diffusing single-molecule experiments.
Figure 2.12 .................................................................................................................................. 41
Images of the scanning (top) and time trajectory (bottom) windows of the
SPCImage.exe software used to acquire fluorescence data.
Figure 2.13 .................................................................................................................................. 42
Image of the vffta64.exe software use to analyze the data acquired via the
SPCImage.exe program.
Chapter 3:
xvi

Figure 3.1 .................................................................................................................................... 49
Cutaway diagram of IR absorption based heating apparatus. Inset shows the absorption
coefficient (α) of water in the vOH = 2←0 overtone band region near 1.45 µm.
Figure 3.2 .................................................................................................................................... 52
Arrhenius analysis of the non-radiative component knrad(T) for temperature- dependent
rhodamine B fluorescence rates. By explicitly measuring kfl = krad + knrad (T) andmodeling knrad(T) as a thermally activated Arrhenius process, one can use TCSPC andthe Arrhenius equation as a calibration curve for temperature.
Figure 3.3 .................................................................................................................................... 52
Van’t Hoff plot of Rln[Keq] vs 1/T for single molecule FRET studies of RNA folding
provides ΔH° and ΔS° for the GAAA tetraloop−receptor tertiary interaction. Note theexcellent agreement between the laser-overtone and stage-based heating techniques.
Figure 3.4 .................................................................................................................................... 54
Time scale for IR-absorption based heating. By monitoring rhodamine B fluorescence
lifetimes, one can directly measure characteristic times associated with laser heating of
the sample. The characteristic heating time is currently limited by the time binning
resolution of the data (< 20 ms), which is slower than, but consistent with, predictions(see section 3.2 for details).
Chapter 4:
Figure 4.1 .................................................................................................................................... 64
Schematic representation of the infrared heating apparatus illustrating the coaxiallyaligned foci of the infrared (red) and excitation (green) lasers within a standard sample
holder (not to scale). The intensity plot depicts the experimentally measured radialintensity profile at the focus of the infrared laser (ωo = 10.04(4) µm).
Figure 4.2 .................................................................................................................................... 66
Fluorescence decay profile for Rhodamine B as a function of temperature. The
linearity of the fluorescence decay over three orders of magnitude attests to the singleexponential behavior of Rhodamine B. The inset represents the Eyring analysis of thetemperature-dependent non-radiative rate constant for Rhodamine B (see section 4.2.1
for details), which is used to determine the temperature rise in the diffraction-limited
confocal volume of the sample.
Figure 4.3 .................................................................................................................................... 68
Experimentally measured steady state temperature rise (ΔT) as a function of (a)incident laser power as well as (b) radial and (c) axial distances from the laser focal
point. The solid red region in (b) represents the radial intensity profile of the infraredlaser beam scaled to the maximum ΔT. Note the clear broadening of the temperature
profile beyond the laser excitation region, which arises largely from thermal diffusionaway from the heating axis.
Figure 4.4 .................................................................................................................................... 71
Real time observation of the infrared heating and cooling behavior, as obtained byxvii

time-correlated single-photon fluorescence measurements of Rhodamine B in theconfocal excitation region. (a) Sample temperature trajectory (black ⊗’s) resulting
from a 50 mW heating pulse with a duration of 500 ms. The solid lines represent multiexponential fits used to calculate the half-maximal times for heating (solid red) andcooling (solid blue). (b) A linear-log plot demonstrating the dependence of coolingtime constant on the heating duration, which suggests that the system achieves a steadystate thermal distribution on the few second time scale.
Figure 4.5 .................................................................................................................................... 74(a) Diagram of the duplex DNA fluorescent construct used for the dissociation
experiments. Green and red stars represent the Cy3 and Cy5 FRET pair used in theexperiments. The solid cyan line corresponds to the PEG18 internal linker within theconstruct to easily allow the construct to form the closed conformation. (b) ALEX
surface image (10 µm × 10 µm) prior to and (c) after exposure to a pulse of infrared
light. White circles represent the mask used to measure survival probability (seesection 4.3.1 for details).
Figure 4.6 .................................................................................................................................... 76
ALEX scanned image (80 µm × 80 µm) of the fluorescent DNA duplex constructs after
heating the center of the image to 72.3(5) °C for 10 sec. Note the clear circular regionof high FRET molecules (red), corresponding to duplex constructs that have undergone
dissociation, while the surrounding low FRET species (green) remain in the duplex
conformation.
Figure 4.7 .................................................................................................................................... 79(a) DNA duplex survival probability data, S(Δt,T), as a function of heating duration, Δt,
at four steady-state heating pulse temperatures. Dashed lines represent least squares fit
to a consecutive first order reaction model, which yields n = 4.1(3) sequential reaction
steps with an assumed identical rate constant. Notice the rapid increase in this rate
constant with increasing temperature, which implies a large endothermic component
the transition state barrier. (b) Eyring transition state analysis of the temperaturedependent rate constant, suggesting a highly endothermic (ΔH‡ = 50(10) kcal/mol) andentropically rewarding (ΔS‡ = 100(20 cal/mol/K) component of the free energy barrier
for each of the fraying events.
Figure 4.8 .................................................................................................................................... 83
Model depicting the “consecutive fraying at both ends” mechanism for duplex
dissociation. The colored bars represent short segments of the 21 bp DNA duplex thatare either associated (green) or dissociated (red). The transparent black numbers
correspond to the relative abundance of each partially dissociated species as predictedby a binomial distribution. After four consecutive fraying events, the single remaining
associated (green) segment of the duplex is sufficiently frayed to undergo a final rapid
dissociation step (see section 4.4 for details).
Chapter 5:
Figure 5.1 .................................................................................................................................... 97
Schematic of the three-piece donor/acceptor label RNA construct. The isolated GAAA

xviii

tetraloop−receptor serves as a model system to investigate the thermodynamics andkinetics of monovalent-facilitated formation of tertiary interactions in structured
RNAs.
Figure 5.2 .................................................................................................................................... 98(a) Sample time trajectories for the model construct. A hidden Markov fit to the FRET
trajectory is used to demonstrate the two-state behavior of folding process. (b)Cumulative distribution function (CDF) of the dwell times demonstrates singleexponential behavior and are used to extract unimolecular rate constants for folding
and unfolding.
Figure 5.3 .................................................................................................................................. 100(a) Na+-dependent kinetics for the formation of the GAAA tetraloop−receptor
interaction. Increasing [Na+] yields a significant enhancement of the folding rate
constant (kfold) and a subtle reduction in the rate constant for the unfolding process(kunfold). (b) The dependence of the free energy on [Na+] and (c) preferential interaction
coefficient analysis for the formation of the tertiary interaction, which yields an ion
pair uptake of 2ΔΓ± = 2.4(1) and no change in free energy at [Na+] = 170(20) mM.
Figure 5.4 .................................................................................................................................. 102(a) van’t Hoff analysis of the tetraloop−receptor interaction at four [Na+]. Increasing[Na+] increases both the ΔH° (decreasing slope) and the ΔS° (increasing verticalintercept) for the formation of the tertiary interaction. (b) Table of thermodynamic
parameters from the van’t Hoff analysis. (c) Bar graph to visually display the change in
ΔH° and ΔS° as a function of ln[aNa+].
Figure 5.5 .................................................................................................................................. 107(a) Transition state theory (TST) analysis of the rate constant associated withformation of the tetraloop−receptor interaction (kfold) as a function of temperature. (b)Table of thermodynamics parameters from the TST analysis. (c) Bar graph depicting
the change in ΔH‡fold and ΔS‡fold at four values of ln[aNa+].
Supplemental Figure S5.1 ....................................................................................................... 110
Schematic of the three-piece donor/acceptor label RNA construct. The poly-U7 linker
is used to determine the effect of the linker on the folding thermodynamics.
Supplemental Figure S5.2 ....................................................................................................... 111(a) van’t Hoff analysis of tetraloop−receptor interaction at four [Na+] with the poly-U7
linker. Increasing [Na+] increases both the ΔH° (decreasing slope) and the ΔS°(increasing vertical intercept) for the formation of the tertiary interaction. (b) Table ofthermodynamic parameters from the van’t Hoff analysis. (c) Visual display of changes
in ΔH° and ΔS° as a function of ln[Na+].
Supplemental Figure S5.3 ....................................................................................................... 111(a) Transition state theory (TST) analysis of the rate constant associated withformation of the tetraloop−receptor interaction (kfold) as a function of temperature withthe poly-U7 linker based construct (b) Table of thermodynamics parameters from thexix

TST analysis (c) Bar graph depicting the change in ΔH‡fold and ΔS‡fold at four values ofln[Na+].
Figure 5.6 .................................................................................................................................. 112
Complete thermodynamics reaction coordinates for the formation of the GAAA
tetraloop−receptor tertiary interaction. The three panels depict the: (a) enthalpy, (b)entropy, and (c) free energy along the reaction coordinate at 180 and 630 mM Na+. The
horizontal distance between states qualitatively represents how “early” or “late” they
are along the reaction coordinate (see Section 5.4 for details).
Figure 5.7 .................................................................................................................................. 117
Structural representation of the free and bound receptor domain. Bases that undergo
significant structural rearrangement upon forming the tetraloop−receptor interaction
are color-coded. Bases of the GAAA tetraloop are not shown to emphasize changes
within the receptor domain (see Section 5.4.3 for details).
Figure 5.8 .................................................................................................................................. 121
Monovalent-identity and folding kinetics for the formation of the GAAA
tetraloop−receptor interaction. Increasing monovalent concentration provides asignificant enhancement of the folding rate constant (kfold) and a subtle reduction in therate constant for the unfolding process (kunfold). No significant difference is observed
between Na+ and K+.
Figure 5.9 .................................................................................................................................. 122(a) van’t Hoff analysis of tetraloop−receptor interaction comparing [Na+] and [K+](b)Eyring analysis of tetraloop−receptor interaction comparing [Na+] and [K+]. (c,d)
Table of thermodynamic parameters from the van’t Hoff and Eyring analyses. As
depicted, folding behavior is largely independent of monovalent cation identity.
Chapter 6:
Figure 6.1 .................................................................................................................................. 133
Free energy diagram representing the effect of TMAO or urea on the stability of theunfolded (U) or folded (F) structures of a nucleic acid. TMAO has a more unfavorable
interaction with the unfolded structure than the folded structure, which results in a net
decrease in ΔG°fold. Urea has a more favorable interaction with the unfolded
conformation than folded conformation, which gives rise to a net increase in ΔG°fold.
Figure 6.2 .................................................................................................................................. 135
Cartoon representation of the two smFRET nucleic acid constructs used throughout the
study. The DNA duplex construct (a and b) is intended to be a model system fornucleic acid secondary structure, where as the TL-TLR construct (c and d) serves as amodel for tertiary interactions in structured RNAs. For both constructs, formation of
the interaction (b and d), referred to as folding, results in an increase in EFRET (a and c).
Figure 6.3 .................................................................................................................................. 137
Representative donor (green) and acceptor (red) fluorescence trajectories for (a) the
TL-TLR and (b) the DNA duplex. Fluorescence data is use to calculate the FRETxx

trajectories (black), which are then analyzed to measure the folding (kfold) and
unfolding (kunfold) rate constants for each of the nucleic acid constructs.
Figure 6.4 .................................................................................................................................. 138
Kinetic origin of osmolyte-influenced nucleic acid folding. The stabilizing osmolytes
TMAO (blue box) increases (a) the folding rate constant (kfold, red) and decreases the(b) unfolding rate constant (kunfold, green) for both the GAAA tetraloop—receptor(triangle) and DNA duplex (circle) constructs. The destabilizing osmolyte, urea,
decreases the (c) folding rate constant (kfold, red) and increases the (d) unfolding rate
constant (kunfold, green) for both constructs.
Figure 6.5 .................................................................................................................................. 139
Osmolyte-dependent folding free energy change for (a) the DNA duplex and (b) the
TL-TLR constructs. For both constructs, addition of TMAO (blue stars) always
stabilizes the folding transitions, where as addition of urea (orange squares)
destabilizes the folding transitions as indicated by the dependence of ΔG° on theosmolyte concentration.
Figure 6.6 .................................................................................................................................. 141
van’t Hoff plots for the osmolyte-influenced folding of both (a) the DNA duplex and(b) the TL-TLR interaction. Solid lines in (a) correspond to a global fit with a common
slope, as discussed in the text, to reduce parameter correlation between the three data
sets: 1 M TMAO (blue stars), 0 M osmolyte (pink circles), and 1 M urea (orange
squares). The solid lines for the TL-TLR interaction (b) are also the result of a global
fit of the 0 M osmolyte and 1 M urea data sets in order to reduce parameter correlation,
while the solid line for the 1M TMAO data set represents an independent fit of thedata. See Table 6.1 for values of the fitted parameters.
Supplemental Figure S6.1 ....................................................................................................... 142
Temperature-dependent kinetic data for the (a) folding and (b) unfolding rate constantsfor the DNA duplex. The data are analyzed using transition state theory. Solid lines incorrespond to a global fit with a common slope, as discussed in the text, to reduceparameter correlation. The large entropic offset in the temperature-dependence of theunfolding rate constants (b) suggests that most of the change in ΔS° arises fromchanges in the entropic barrier to unfolding (ΔS‡unfold). The data table (c) contains
values for the transition state theory analysis of the temperature-dependent kinetic data
for the DNA duplex. Fitting methods are identical to those used in the van’t Hoffanalysis (Figure 6.6), which is discussed in section 6.3.4.
Supplemental Figure S6.2 ....................................................................................................... 144
Temperature-dependent kinetic data for the (a) folding and (b) unfolding rate constantsfor the TL-TLR. The data are analyzed using transition state theory. Solid lines for 1 M
urea (orange squares) and 0 M osmolyte (pink circles) correspond to a global fit with acommon slope, as discussed in the text, to reduce parameter correlation. The large
entropic offset for 1 M urea in the temperature-dependence of the folding rate constant(a) suggests that most of the change in ΔS° arises from changes in the entropic barrier
to folding (ΔS‡fold). The distinct thermodynamic behavior of TMAO- facilitatedxxiformation of the TL-TLR interaction is discussed in the text. The data table (c)contains values for the transition state theory analysis of the temperature-dependent
kinetic data for the DNA duplex. Fitting methods are identical to those used in the
van’t Hoff analysis (Figure 6.6), which is discussed in section 6.3.4.
Figure 6.7 .................................................................................................................................. 155
Schematic representation of the inverted confocal fluorescence microscope used toacquire smFRET trajectories. Individual molecules are immobilized to the glass
coverslip using standard biotin-streptavidin chemistry. Excitation and emission isachieved using standard epifluorescence techniques. Fluorescent photons are focusedthrough a confocal pinhole before being separated by color and polarization anddirected onto 4 single photon avalanche photodiodes. Upon detection, the arrival times
of individual photons are recorded and sent to a time-correlated single-photon counting(TCSPC) router where they are used to generate trajectories of donor and acceptor
fluorescence.
Chapter 7:
Figure 7.1 .................................................................................................................................. 163
Details of the human telomerase RNA pseudoknot a) Secondary structure diagrams ofthe minimal pseudoknot motif in a partially unfolded (hairpin) conformation and afolded (pseudoknot) conformation. The GC!AG mutation at the orange nucleotides in
the psuedoknot region corresponds to the genetic disorder dyskeratosis congenita.
Green (Cy3) and red (Cy5) stars indicate the location of two fluorophores, where sizeof the stars reflect FRET efficiency between the two dyes and the black vertical bar
represents the ligation site. b) Model solution structure of the minimal pseudoknot
motif, with the same color scheme as the secondary structure diagram. c) Schematicequilibrium between the pseudoknot and hairpin conformations.
Figure 7.2 .................................................................................................................................. 165
Freely diffusing FRET histograms of a) the wild-type human telomerase RNA
pseudoknot and b) the dyskeratosis congenita mutant of the human telomerase RNA
pseudoknot. The urea titrations demonstrate strong shifts in the pseudoknot-hairpin
equilibrium behavior for both constructs.
Supplemental Figure S7.1 ....................................................................................................... 167a) Details of the alternative labeling scheme for the wild-type human telomerase RNA
pseudoknot secondary structure. The minimal pseudoknot motif is in a partiallyunfolded (hairpin) conformation and a folded (pseudoknot) conformation. The green(Cy3) and red (Cy5) stars indicate the location of the two fluorophores, while the sizeof the stars represents the FRET efficiency between the two dyes. The grey vertical bar
represents the location of the ligation site (see Section 7.5.1 for details) b) Freely
diffusing FRET histograms of alternate wild-type human telomerase RNA pseudoknot.
The nearly identical FRET values at high and low urea concentrations confirm that: (i)the P2 region of the pseudoknot remains intact under all experimental conditions and(ii) the dynamics associated with the conventional constructs results fromformation/disruption of the P3 region of the pseudoknot.

xxii

Figure 7.3 .................................................................................................................................. 168
Urea dependence of the free energy change for forming the pseudoknot from thehairpin conformation for the wild-type human telomerase RNA pseudoknot (grey) andthe dyskeratosis congenita mutant (orange) of the human telomerase RNA pseudoknot.
Linear extrapolation to 0 M urea yields the 4.5(3) kcal/mol differential folding freeenergy between the WT and mutant pseudoknot under non-denaturing conditions.
Figure 7.4 .................................................................................................................................. 169
Images of surface immobilized molecules for the wild-type a,b) and dyskeratosis
congenita c,d) pseudoknots. Each diffraction limited fluorescence spot represents thelocation of an individual dually labeled RNA molecule. The urea dependence of theimages yields kinetic information about the two different pseudoknot constructs (seeSection 7.2.2 for details).
Figure 7.5 .................................................................................................................................. 170
Surface immobilized folding and unfolding kinetics for the a) wild-type pseudoknot
and c) the mutant human telomerase RNA pseudoknots under denaturing and nondenaturing conditions, respectively. Rate constants are determined from exponential
fits to cumulative distribution plots of dwell times for the b) wild-type and d)dyskeratosis congenita pseudoknots.
Figure 7.6 .................................................................................................................................. 172
Log-linear plot of the urea-dependent folding and unfolding kinetics for the a) wildtype and b) dyskeratosis congenita pseudoknots. For both constructs, the unfolding rate
constant is substantially more sensitive to addition of urea than the folding rate
constant, which suggests that most of the change in base solvent accessible surface
area (SASA) occurs after formation of the transition state. Lines indicate trends and are
not actual fits to the data.
Figure 7.7 .................................................................................................................................. 175
Single-molecule van’t Hoff analyses. a) A plot of the temperature-dependence of theequilibrium constant of the wild-type human telomerase RNA pseudoknot at three urea
concentrations. The identical slopes indicate a urea-independent folding enthalpy,
while the constant vertical offset indicates a urea-dependent entropy. b) A plot of theurea-dependent entropy suggests a linear dependence of folding entropy on urea
concentration. c) van’t Hoff analysis of the mutant pseudoknot under non-denaturingconditions. All entropy and enthalpy changes can be found in Table 7.2.
Figure 7.8 .................................................................................................................................. 178
Free energy landscape for formation of the pseudoknot from the hairpin conformation.
Secondary structure diagrams for the wild-type human telomerase RNA pseudoknot
are shown, including the predicted structure of the transition state where only a few
nucleotides from the P3 region of the pseudoknot have formed (see Section 7.3.1 fordetails).
Figure 7.9 .................................................................................................................................. 186
Single-molecule fluorescence microscope. Diagram of the experimental setup and dataxxiii

collection (see Section 7.5.2 for details).
Figure 7.10 ................................................................................................................................ 188
Freely diffusing experiments. a) Schematic representation of single-molecules freely
diffusing in and around the overlapping excitation volumes of the red and green lasers(see Materials and Methods for details). b) Freely diffusing data of the humantelomerase RNA pseudoknot obtained using ALEX methods, with the four possible
outcomes for the fluorescent molecules. ALEX filtering permits events from duallylabeled donor-acceptor molecules to be selectively considered for data analysis.
Chapter 8:
Figure 8.1 .................................................................................................................................. 198
Schematic representation of the a) secondary and b) tertiary structure of theenv8HyCbl riboswitch, complete with the Cy3 (green star), Cy5 (red star), and biotin(blue rectangle) synthetic moieties. The loops, L5 (red) and L13 (green) form aregulatory RNA kissing-loop (KL), which contains the purine-rich ribosome bindingsite (shadowed green nucleotides). Binding of hydroxocobalamin (bronze spheres,
HyCbl) facilitates formation of the KL, which decreases the distance between the Cy3(donor) and Cy5 (acceptor) fluorophores resulting in more efficient energy transfer(EFRET).
Figure 8.2 .................................................................................................................................. 199
Four-state kinetic model for the env8HyCbl riboswitch represented a) graphically andb) symbolically. The four macroscopic conformations are linked by three coupledequilibria: (i) KL docking in the absence of ligand, (ii) ligand binding in the undocked
conformation, and (iii) KL docking in the presence of ligand. Formation of the KL
decreases the inter-dye distance, resulting in more efficient fluorescence resonanceenergy transfer (EFRET). Ligand binding quenches Cy3, which decreases the total
fluorescence of Cy3 and Cy5 and can be monitored independent of KL
docking/undocking.
Figure 8.3 .................................................................................................................................. 202
Freely diffusing smFRET experiments depict the Mg2+-dependence of the KL docking
equilibrium in the a) absence and b) presence of 1000 nM HyCbl. c) The presence ofHyCbl (bronze circles) decreases the concentration of divalent cations (i.e., [Mg2+]1⁄2)
required for FD = 0.5.
Supplemental Figure S8.1 ....................................................................................................... 204
Results of freely diffusing experiments conducted on the L5-mutant env8HyCbl
construct. The L5-mutation (inset, underlined nucleotides) inhibits formation of the KL
interaction, which prevents occlusion of the ribosome binding site (inset, shadowed
nucleotides).
Figure 8.4 .................................................................................................................................. 206
Time-dependent ensemble fluorometry. a) HyCbl binding quenches the fluorescence ofCy3 (Supplementary Figure S8.2). b) Elevated concentrations of ligand increases therate constant for establishing equilibrium. c) A linear-fit to the [HyCbl]-dependence ofxxivthis rate constant is used to measure the apparent ligand binding (appkbind) anddissociation (appkdiss) rate constants.
Supplemental Figure S8.2 ....................................................................................................... 207
The fluorescence lifetime (τfluor) of Cy 3 as determined from surface-immobilized
riboswitches bound to (light green triangles) or free of (dark green circles)
hydroxocobalamin (HyCbl).
Figure 8.5 .................................................................................................................................. 210
Freely diffusing single-molecule burst titrations of hydroxocobalamin (HyCbl) at
various concentrations of MgCl2. a) At 0 mM Mg2+, addition of ligand does not
significantly influence distribution of FRET values associated with the L5-L13
regulatory switch. b) At intermediate concentrations of Mg2+(e.g., 0.5, 1.0, and 2.5mM), addition of HyCbl does significantly alter the FRET distributions. c) 20 mM
Mg2+ is sufficient to completely form the L5-L13 KL interaction, irrespective of[HyCbl].
Figure 8.6 .................................................................................................................................. 211
Experimental validation of the four-state kinetic model for the env8HyCbl riboswitch.
The experimentally determined fractional occupancy of the docked conformation (FD,colored circles) is well described by the steady-state solution (dotted line) to the fourstate model (Figure 8.2) over a wide range of Mg2+ and HyCbl concentrations.
Figure 8.7 .................................................................................................................................. 212
Kissing-loop docking kinetics in the absence of bound ligand. Representative a)
fluorescence and b) FRET time-trace from a surface immobilized env8HyCbl
riboswitch. c) Rate constants are determined using both a dwell time analysis and a 2state Bayesian maximum- likelihood model (orange line). d) Kinetic parameters areused to calculate the fractional occupancy of the docked conformation in the absenceof HyCbl (FDfree), which reveals that the surface immobilized results are in accordancewith the freely diffusing experiments.
Figure 8.7 .................................................................................................................................. 214
Surface immobilized KL docking kinetics in the presence of hydroxocobalamin(HyCbl). a) Representative fluorescence trajectory from a surface immobilized
molecule in the presence of 2.5 nM HyCbl. While in the undocked conformation,
ligand binding (arrow) significantly diminishes the fluorescence intensity, which
returns to normal after dissociation. b) When the ligand is not bound, the docking
kinetics of the regulatory KL are experimentally indistinguishable from those in theabsence of any HyCbl (Figure 8.7b). c) When the ligand is bound, a maximumlikelihood 2-state model (black line) is used to quantify the dwell times associated withthe subtle fluctuations in acceptor fluorescence resulting from KL docking andundocking. A comparison between the two KL docking equilibria reveals that HyCbl
binding significantly decreases the undocking rate constant (kundock).

xxv

Supplemental Figure S8.3 ....................................................................................................... 215
Representative surface immobilized trajectories revealing the HyCbl-induced decreasein fluorescence intensity of both Cy3 (green squares) and Cy5 (pink circles). Inspectionof trajectories reveals that HyCbl binding events (black arrows) can only occur when
the RNA is in the undocked (low EFRET) conformation. This is consistent with theassumptions of our 4-state kinetic model, whereby formation of the L5-L13 KLinteractions prevents HyCbl binding/dissociation events. Only trajectories that exhibit
clean fluctuations in acceptor fluorescence after a decrease in total fluorescence areconsidered for analysis. This constraint excludes molecules that have undergone otherphotophysical processes resulting in a decrease in fluorescence (e.g., blinking or
bleaching).
Figure 8.9 .................................................................................................................................. 217
Cell-based regulation of gene expression via the env8HyCbl riboswitch. a) Schematic
diagram relating the four-state kinetic model to the regulation of gene expression in vivo. b)
Histograms comparing the normalized fluorescence and relative GFPuv expression of cell
cultures grown in the presence (bronze bars) or absence (grey bars) of 5000 nM HyCbl.

xxvi

Chapter 1

Introduction

This chapter serves as a broad and general overview of the scientific experiments
contained within this thesis. Chapter 2 provides a thorough and technical description of all of theexperimental procedures required for each of the research topics contained herein. A morecomprehensive and scientific summary of these projects can be found in their respective chapters(i.e., Chapter 3-8).
1.1

Nucleic Acids
Some of the most essential components of all living organisms are nucleic acids becauseof the fundamental role they play in the processes of life. Originally discovered in the 1860s (1),they are polymeric biomolecules comprised of chemically linked building blocks, called
nucleotides (Figure 1.1). Each nucleotide consists of three components: (i) a cyclic 5-carbon
sugar whose 5! oxygen and 1! carbon atoms are attached to (ii) a phosphate group and (iii) a
nitrogenous base, respectively. These building blocks are linked together, forming a polymericchain, by phosphodiester bonds between the sugar and phosphate moieties. One of the
distinguishing characteristics among the various classes of nucleic acids is the type of sugar
molecule. The presence or absence of a hydroxyl group (-OH) on the 2! carbon of the ribose
sugar differentiates between ribonucleotides and deoxyribonucleotides. Covalently linked
together, these nucleotides form the two major types of nucleic acids—ribonucleic acid (RNA)and deoxyribonucleic acid (DNA). The vast majority of nucleic acids consist of sequentially1linked nucleotides, each with one-of-four different nitrogenous bases—(i) Guanine (G), (ii)
Cytosine, (iii) Adenine (A) and (iv) either Thymine (T) in DNA or Uracil (U) in RNA.

Figure 1.1 Structural components of a nucleic acid. (a) Hydrogen bonding pattern between pairsof guanine and cytosine nucleobases. (b) Hydrogen bonding pattern between pairs of adenine
and thymine nucleobases in DNA. (c) Hydrogen bonding pattern between pairs of adenine anduracil nucleobases in RNA. (d) Schematic representation of the phosphate, sugar, and baseconstituents in the context of duplex deoxyribonucleic acid.
The specific sequential arrangement of nucleotides in DNA, typically written starting from theunlinked 5! end of the polymer, constitutes the genetic code of all living organisms. The ability
for DNA to compose the genetic code, or genome, is one of the most well known functions of
nucleic acids(2). Living organisms are continually interpreting this warehouse of information inorder to carry out the plethora of chemical tasks essential for life. Historically, it was thought that2

execution of a particular biological task is accomplished by creating an RNA-based copy
(messenger RNA, mRNA) of a specific segment of the genome—a process called
transcription(3). The function of this RNA molecule was to deliver the genetic message to a large
biological machine, called the ribosome, which would construct the encoded protein—a process
called translation. This linear transfer of genetic information is commonly referred to as theCentral Dogma of Molecular Biology (Figure 1.2).

Figure 1.2 Central Dogma of Molecular Biology. Schematic representation of the linear transfer
of genetic information in a cell. Pointed and blunted arrows represent some of the many waysthat RNA can influence the processing of this genetic information.
For a number of years, it was thought that proteins were the molecular agents responsible forcarrying out most biological tasks. However, over the past 60 years it has become increasingly
apparent that this represents an oversimplified view of life. Nucleic acids, specifically RNAs, arecapable of performing many biological processes(4).
As is commonly the case throughout all of biology, the function of a molecule is strongly
influenced by its molecular structure, and nucleic acids are no exception. The functional
diversity of nucleic acids arises almost completely from the diversity of nucleotide sequences, or
primary (1°) structures available to nucleic acids. Nucleic acid secondary (2°) structure arises
predominantly from intra- or intermolecular interactions among individual nucleotides. These 2°
structural interactions are held together by favorable contacts between pairs of nucleotides.
3

Specifically, the nucleotides G and C are able to pair together in a precise orientation to form
three favorable hydrogen-bonds, while the nucleotides A and T (or A and U in RNA) are able topair and form two hydrogen-bonds (Figure 1.1). One of the results of base pairing is that two
nucleic acids with complementary sequences, when oriented appropriately, are able to favorablyinteract with one another. This was beautifully demonstrated in 1953 with the discovery of the
double helical structure of duplex (i.e., double stranded) DNA(5).
As opposed to the intermolecular 2° structure of duplex DNA (comprising all living
genomes), the single stranded nature of RNA results in primarily intramolecular 2° structures.
For any RNA molecule that is not perfectly self-complementary, intramolecular base pairing will
result in structures that have regions of unpaired nucleotides that join together the double helicalregions of paired nucleotides to form 2° structural elements like loops and junctions. One
advantage of the unpaired junctions commonly found in RNAs is that they are more
conformationally flexible than the paired regions. This allows formation of additional favorable
intramolecular interactions (sometimes canonical base pairs) between individual 2° structuralelements like loops. The interactions among elements of 2° structure result in the formation of
tertiary (3°) structural elements (Figure 1.3). A single RNA polymer can simultaneously form
many different 2° structural elements that can associate with each other in a complex network of
3°interactions. For an RNA of 130 nucleotides, there are >1078 unique 1° structures (e.g.,
sequences). A single copy of each of the unique sequences would have a combined mass that isnearly identical to that of the observable universe (1.6×1055 g (6)). Some of these sequences
would have the potential to form 2° and 3° interactions, each of which would result in a moleculewith a slightly different molecular conformation. It is this diverse array of intricate three-4dimensional structures that gives rise to localized, chemically diverse environments that have thepotential to perform a myriad of useful biological functions.

Figure 1.3 Hierarchical folding of RNA. Example depicts the major structural transitions
associated with the formation of a tRNA molecule. Colors correspond to base paired regions
within the oligonucleotide.
1.2

Nucleic Acid Conformational Transitions
It is important to mention that for all chemical reactions there is a characteristic time-scale associated with how long a particular species will exist before undergoing a chemicaltransformation. There are two general classes of nucleic acid chemical transformations, orconformational transitions, that are relevant to this thesis. The first is formation (or disruption) of
2° structures from 1°structures (Figure 1.3). Since these conformational transitions give rise to,or break apart, paired regions of nucleotides with a double helical structure, they are often
referred to as duplex association/dissociation events. The second class of conformationaltransitions that pertain to this thesis is the formation (or disruption) of 3° structures from2°structures (Figure 1.3). Because these types of transitions result from interactions between
2°structural elements (e.g., loops) they will be referred to as either loop docking/undocking
events or sometimes more generally as folding/unfolding events. Of course, the time-scales
5associated with these structural transitions also influence the ability for the nucleic acid to have auseful biological function. Stated more explicitly, the biological function of a nucleic acid is
influenced not only by its structure, but also by the dynamics of the relevant structure.
The five nucleic acid conformational transitions described in this thesis are as follows(Figure 1.4):

Figure 1.4 Cartoon graphics depicting the numerous oligonucleotide conformation transitions
discussed in the chapters of this thesis.
(i) a 21 bp DNA duplex(7) that serves as a simple model system to understand the fundamentals
of nucleic acid duplex association and dissociation, which has relevance in the field of
microRNAs, (ii) an 8 bp DNA duplex inspired by the telomereic repeat sequence in the humangenome which also serves as a model to better understand 2° structure formation(8) (iii) a
ubiquitous 3° structural interaction from an RNA that aids in the maturation of pre-mRNAs—the6

GAAA tetraloop-receptor from the Tetrahymena ribozyme(9), (iv) an isolated 3° structuralelement from an RNA that helps prevent damage from occurring at the ends of linear genomic
DNA—the human telomerase RNA pseudoknot(10), and (v) an entirely functional element ofsome bacterial mRNAs with a 3° interaction that prevents translation by the ribosome—the env8
vitamin-B12 riboswitch.
1.3

Cosolute-Influenced Nucleic Acid Conformational Transitions
One common feature of all nucleic acid conformational transitions, either in vitro or invivo, is that the favorability (or stability) of the interactions and/or the time-scales associatedwith these structural changes are influenced by the presence of other cosolutes in solution(11).
The most common cosolutes associated with nucleic acid conformational transitions are thebiologically relevant alkali and alkaline earth salts of NaCl, KCl, CaCl2 and MgCl2. In aqueous
solutions, these salts readily dissociate into their ionic components of either monovalent (i.e., K+
or Na+) or divalent (i.e., Ca2+ or Mg2+) cations and monovalent anions (i.e., Cl-). As cosolutes ina solution with dissolved ions, nucleic acids are able to interact with these charged species.
Specifically, the negative charge associated with each phosphate in the backbone of a nucleic
acid excludes proximal anions, which reduces the local concentration (12). The opposite is true
for positively charged anions, which are accumulated near the nucleic acids, thus increasing the
local concentration. This effectively neutralizes the high negative charge density of a nucleic
acid. When nucleic acids undergo transitions to higher-order conformations, they tend to become
more compact and as a result the molecule will have an increased negative charge density(13).
Accordingly, additional cations or anions are accumulated or excluded from around the nucleic
acid, respectively, so as to compensate for the increased negative charge density. Because of thegreater positive charge density associated with higher valance cations (e.g., Mg2+ and Ca2+), a7

smaller number of them need to be accumulated in order to accomplish the same degree ofcharge neutralization. This is partially responsible for lowering the cation concentrations
required for half-maximal folding (i.e., [Mn+]1/2) from ~ 100 mM for monovalent cations to ~ 1
mM for divalent cations and ~0.1 mM for trivalent cations(11). The influence of metal cations on
nucleic acid conformational transitions is the topic of Chapter 5 in this thesis and covered
extensively in an entire thesis from a former graduate student in the lab—Dr. Julie Fiore. Briefly,
Chapter 5 describes how experiments were performed to obtain a better understanding of how
monovalent cations facilitate formation of the GAAA tetraloop-receptor interaction from the
Tetrahymena ribozyme(14).
Although metal cations are some of the most common cosolutes associated with nucleic
acid conformational transitions, there are many others, including non-ionic organic molecules.
The most commonly used cosolute to destabilize nucleic acid structures is the non-ionic organic
osmolyte urea(15)(Figure 1.5). Osmolytes are highly soluble chemical compounds that can beused by cells to control osmotic pressure(16). High concentrations of urea are known to hinder
formation of both 2° and 3° structures in both DNA and RNA(17). This is due to favorable
dipole-driven base stacking and hydrogen bonding interactions between urea and the nucleobases(18) that can out compete the corresponding water-nucleobase interactions. Accordingly, urea
tends to accumulate around the solvent accessible surface area (SASA) of these nitrogenous
bases, which stabilizes any given nucleic acid structure. However, nucleic acid conformationaltransitions (e.g., duplex association) frequently decrease the amount of base SASA leading to theunfavorable expulsion of urea from the buried surfaces. This phenomenon results in a lesseffective stabilization of higher-order nucleic acid structures (i.e., 1° structure > 2° structure > 3°8

structure). As a result, urea has a net destabilizing effect on more compact and folded
conformations, relative to the more voluminous unfolded conformations.

Figure 1.5 Chemical structures of two osmolytes known to significantly influence theconformational transitions of nucleic acids.
Surprisingly, there are other osmolytes, for example trimethylamine oxide (TMAO), thatcan stabilize nucleic acid conformational transitions (Figure 1.5). TMAO interacts unfavorablywith the phosphate SASA of nucleic acids (19), and, although the details of this unfavorable
interaction are not entirely understood, it is also likely related to the large dipole-moment of themolecule. However, unlike urea, the three methyl groups sterically impair dipole-driven
interactions between TMAO and the phosphates, rendering this osmolyte as a less effective
solvent than water. However, as mentioned earlier, conformational transitions responsible for theformation of 2° and 3° structures (e.g., loop docking) decrease the amount of SASA. Therefore,
when an RNA becomes more compact, TMAO is ejected from the buried surfaces, which isenergetically rewarding. As a result, TMAO is less detrimental to higher-order, more compact
nucleic acid structures, leading to a net stabilization. The influence of both osmolytes—TMAO

9

and urea—on nucleic acid conformational transitions represents the research topic of Chapter 6in this thesis.
Additionally, the destabilizing ability of urea is used as a biochemical tool in Chapter 7 ofthis thesis to study how a two-nucleotide substitution mutation associated with a genetic disorderinfluences an RNA conformational transition required for proper biological activity of an
enzyme—telomerase—that is highly associated with cancer(10). There are many other non-ionicorganic molecules (e.g., molecular crowding agents like polyethelyne glycol and viscous agentslike glycerol) that non-specifically influence nucleic acid conformational transitions. The reportsof these investigations have been published in peer-reviewed journals in conjunction withanother Nesbitt group member, Nick Dupuis, as first author and thus are not included in thisthesis.
Contrary to the non-specific effects of all of the cosolutes described above, certain
nucleic acid conformational transitions are known to be particularly sensitive to specificcosolutes. The specificity of the interaction is achieved by precise intermolecular contacts
between discrete functional groups on the cosolute and nucleic acid. Because of the specificity
associated with this class of cosolute-nucleic acid interactions, the cosolutes are often referred toas biochemical ligands, in that they are substances that complex with, or are bound by,biomolecules to produce a chemical response. Some nucleic acids that contain these cosolutespecific conformational transitions have been employed by nature to act as molecular switches
capable of changing their biochemical function upon ligand binding. This switch of biochemicalfunction is most frequently accomplished by ligand-induced conformational transitions. Some of
the most well known examples of ligand-induced conformational transitions in nucleic acids arefound in riboswitches(20, 21). Riboswitches are structural elements genetically encoded at the10beginning of mRNAs that modulate expression of a downstream gene via ligand-dependent
conformational changes. The modulation of gene expression can occur by prematurely
terminating the transcription of the mRNA, or by an inability of the ribosome to initiate
translation of the mRNA. Chapter 8 in this thesis describes the study of a translational riboswitch
that, upon binding vitamin B12, prevents expression of downstream genes.
This thesis covers a number of cosolute-influenced nucleic acid conformationaltransitions. These reports vary both in terms of the type of conformational transition investigated
and the identity of the cosolutes. As a compilation of six scientific publications, this document
provides a detailed review of the numerous ways in which these conformational transitions are
influenced by the presence of aqueous cosolutes.
1.4

Single-Molecule Fluorescence
Because of the small size of molecules, most biochemical experiments are performed at

the ensemble level by studying billions of molecules simultaneously. These types of experiments
have been extremely useful for unveiling numerous details about a number of cosoluteinfluenced nucleic acid conformational transitions. For years, temperature-dependent
ultraviolet/visible spectroscopy and circular dichroism have been used to investigate the stabilityof various nucleic acids in the presence of a diverse array of cosolutes. However, theseexperiments are unable to provide detailed information about the time-scales (i.e., kinetics) of theconformational transitions because the billions of molecules are all changing conformations at
random times. In order for ensemble techniques to obtain useful kinetic information, themolecules must be synchronized so that all of the nucleic acids undergo conformationaltransitions at the same time. Molecular synchronization is most frequently accomplished bywatching a sample return to equilibrium after rapid heating/cooling or in the case of cosolute-11

influenced conformational transitions, rapid mixing. These synchronized experiments often
require various assumptions about which processes are fast, relative to others, in order to yield
meaningful kinetic information. Another assumption that is intrinsic to ensemble experiments isthat the billions of molecules in an experiment are identical, and thus statistically behave exactly
the same. This set of constraints can be nullified by watching individual molecules undergo
discrete chemical transformations—a concept frequently quoted as being “every chemist’s
dream.”The path towards realization of this dream started nearly a quarter-century ago with theobservation of absorption(22, 23) and fluorescence(24) from single molecules in a cryogenic
crystal. Over the past 25 years, single-molecule fluorescence techniques have greatly improved
due to more powerful lasers, better microscope objectives, higher efficiency detectors, andbrighter fluorophores. These advancements have made direct observation of single moleculesmuch more attainable.
Of the many fluorescence methods that can be extended to the single-molecule level, thisthesis exploits the power of Förster Resonance Energy Transfer (FRET)(25-27) to observe single
nucleic acids undergo cosolute-influenced conformational transitions in real-time. In 1948,
Theodore Förster was the first to correctly define the physics behind the process of FRET(28),which describes the transfer of energy between two molecular species that absorb light (i.e.,
chromophores). Most commonly, and of particular relevance to this thesis, these chromophores
also emit light (i.e., fluorophores). After absorbing a photon of a specific color, the electronicarrangement of a ground state molecule is altered to a higher-energy excited configuration. In theabsence of an acceptor fluorophore, the excited donor will relax back down to the ground stateelectronic configuration and, in the process, potentially give off a photon(29). However, in the12presence of an acceptor fluorophore, the excited donor can, instead, transfer the excess energy
associated with this excited electronic configuration to an acceptor via near-field nonradiative
dipole-dipole coupling. After an energy transfer event, the newly-excited acceptor fluorophore
will, just like the donor, relax back down to the ground state electronic arrangement andpotentially emit a photon.

Figure 1.6 Fluorescence Resonance Energy Transfer (FRET). An excited donor fluorophore(green) can transfer energy to a nearby acceptor fluorophore (red) in a distance-dependent
process. When the donor and acceptor are proximal, energy transfer is efficient; when they are
distal, energy transfer is poor.
The efficiency of FRET (EFRET) can readily be determined by observing the ratio ofacceptor fluorescence to total fluorescence (i.e., donor plus acceptor). The physics of this energy
transfer process dictate that the EFRET is inversely related to the sixth-power of the distance
between the donor and acceptor fluorophores. The effective distance scale for FRET depends on
the spectral properties of the fluorophores and ranges between 10-100 Å(30). Conveniently, thisis also the distance scale associated with the change in molecular dimensions for many nucleic
13

acid conformational transitions. For any FRET fluorophore pair (i.e., donor-acceptor pair),conformational transitions that decrease the donor-acceptor distance result in an increase in
EFRET (Figure 1.6), as determined by a decrease in donor fluorescence and a corresponding
increase in acceptor fluorescence.
My general strategy for all of the single-molecules studies described within this thesis
can be summarized using the following three steps: (i) First, donor and acceptor fluorophores
must cleverly and nonperturbatively attach to the nucleic acid of interest(31). (ii) Then, a singlemolecule fluorescence microscope with a high numerical aperture objective can be used to excite
and collect the fluorescence from single donor and acceptor labeled nucleic acid. When
combined together, single-molecule fluorescence techniques and FRET (smFRET) make itpossible to watch the donor-acceptor distance of individual nucleic acids change, as a result ofconformation transitions, in real-time. (iii) Finally, cosolutes can easily be added into theexperimental solutions to readily interrogate the influence of these species on the kinetics anddynamics of nucleic acid conformational transitions.
Although smFRET is a useful experimental technique to observe the kinetics ofconformational transitions in nucleic acids, it can be even more informative when combined withthe ability to precisely control the temperature at which experiments are performed(7, 14, 32-34).
This provides the ability to understand various thermodynamic aspects of nucleic acidconformational transitions. This topic is the research topic of Chapter 3 in this thesis, which
describes the development of a specialized heating technique using focused infrared laser
light(34). Furthermore, Chapter 4 provides a detailed characterization of the aforementioned
heating technique and demonstrates how it can be utilized to probe the dissociation of a 21 basepair DNA double helix(7).

14In conclusion, this document describes a highly interdisciplinary collection of rigorousand quantitative physical chemistry experiments performed on a diverse array of nucleic acids.
These single-molecule investigations explore the effects of various different aqueous cosolutes
on the conformational transitions associated with these biomolecules. The results of theseundertakings provide a more fundamental understanding of the numerous scientific concepts
related to each of the six chapters. As a whole, the conclusions contain within this body of work
provide the scientific community with a holistic foundation for understanding cosoluteinfluenced conformational transitions that can be expanded to include complex solutions such as
those associated with living cells. One day, these fundaments idea might influence the way weaddress pressing societal issues, particularly those related to human health and wellbeing.

15

1.5

References

1.

Dahm, R. Discovering DNA: Friedrich Miescher and the early years of nucleic acidresearch, Hum. Genet., 2008, 122, 565-581.

2.

Voet, D.; J. G. Voet (2011) Biochemistry, 4th ed., John Wiley & Sons, Hoboken, NJ.

3.

Crick, F. Central dogma of molecular biology, Nature, 1970, 227, 561-563.

4.

Silverman, S. K. (2006) Nucleic acid switches and sensors, Landes
Bioscience/Eurekah.com and Springer Science+Business Media, Georgetown, TX andNew York, NY.

5.

Watson, J. D.; F. H. Crick. Molecular structure of nucleic acids: a structure fordeoxyribose nucleic acid, Nature, 1953, 171, 737-738.

6.

Carvalho, J. C. Derivation of the mass of the observable universe, Int. J. Theor. Phys.,
1995, 34, 2507-2509.

7.

Holmstrom, E. D.; N. F. Dupuis; D. J. Nesbitt. Pulsed IR heating studies of singlemolecule DNA duplex dissociation kinetics and thermodynamics, Biophys. J., 2014, 106,220-231.

8.

Dupuis, N. F.; E. D. Holmstrom; D. J. Nesbitt. Single-Molecule Kinetics Reveal CationPromoted DNA Duplex Formation Through Ordering of Single-Stranded Helices,
Biophys. J., 2013, 105, 756-766.

9.

Hodak, J. H.; C. D. Downey; J. L. Fiore; A. Pardi; D. J. Nesbitt. Docking kinetics andequilibrium of a GAAA tetraloop-receptor motif probed by single-molecule FRET, Proc.
Natl. Acad. Sci. USA, 2005, 102, 10505–10510.

10.

Holmstrom, E. D.; D. J. Nesbitt. Single Molecule FRET Studies of the Human
Telomerase RNA Pseudoknot: Temperature/Urea Dependent Folding Kinetics andThermodynamics, J. Phys. Chem. B., 2014.

11.

Fiore, J. L.; E. D. Holmstrom; L. R. Fiegland; J. H. Hodak; D. J. Nesbitt. The Role ofCounterion Valence and Size in GAAA Tetraloop-Receptor Docking/Undocking
Kinetics, J. Mol. Biol., 2012.

12.

Draper, D. E.; D. Grilley; A. M. Soto. Ions and RNA folding, Annu. Rev. Biophys.
Biomolec. Struct., 2005, 34, 221–243.

13.

Leipply, D.; D. Lambert; D. E. Draper. Ion-RNA interactions: Thermodynamic analysisof the effects of mono- and divalent ions on RNA conformational equilibria, Methods
Enzymol., 2009, 469, 433-463.

14.

Holmstrom, E. D.; J. L. Fiore; D. J. Nesbitt. Thermodynamic origins of monovalentfacilitated RNA folding, Biochemistry, 2012, 51, 3732-3743.

16

15.

Shelton, V. M.; T. R. Sosnick; T. Pan. Applicability of Urea in the Thermodynamic
Analysis of Secondary and Tertiary RNA Folding, Biochemistry, 1999, 38, 16831-16839.

16.

Lambert, D.; D. E. Draper. Effects of Osmolytes on RNA Secondary and Tertiary
Structure Stabilities and RNA-Mg2+ Interactions, J. Mol. Biol., 2007, 370, 993-1005.

17.

Lambert, D.; D. E. Draper. Denaturation of RNA secondary and tertiary structure byurea: simple unfolded state models and free energy parameters account for measured mvalues, Biochemistry, 2012, 51, 9014-9026.

18.

Priyakumar, U. D.; C. Hyeon; D. Thirumalai; A. D. MacKerell. Urea Destabilizes RNA
by Forming Stacking Interactions and Multiple Hydrogen Bonds with Nucleic Acid
Bases, J. Am. Chem. Soc., 2009, 131, 17759-17761.

19.

Lambert, D.; D. Leipply; D. E. Draper. The osmolyte TMAO stabilizes native RNA
tertiary structures in the absence of Mg2+: Evidence for a large barrier to folding from
phosphate dehydration, J. Mol. Biol., 2010, 404, 138-157.

20.

Breaker, R. R. Riboswitches and the RNA world, Cold Spring Harb. Perspect. Biol.,2012, 4.

21.

Garst, A. D.; A. L. Edwards; R. T. Batey. Riboswitches: structures and mechanisms, Cold
Spring Harb. Perspect. Biol., 2011, 3.

22.

Kador, L.; D. E. Horne; W. E. Moerner. Optical-Detection and Probing of Single Dopant
Molecules of Pentacene in a P-Terphenyl Host Crystal by Means of AbsorptionSpectroscopy, J. Phys. Chem., 1990, 94, 1237-1248.

23.

Moerner, W. E.; L. Kador. Optical-Detection and Spectroscopy of Single Molecules in a
Solid, Phys. Rev. Lett., 1989, 62, 2535-2538.

24.

Orrit, M.; J. Bernard. Single pentacene molecules detected by fluorescence excitation in ap-terphenyl crystal, Phys. Rev. Lett., 1990, 65, 2716-2719.

25.

Joo, C.; H. Balci; Y. Ishitsuka; C. Buranachai; T. Ha. Advances in single-molecule
fluorescence methods for molecular biology, Annu. Rev. Biochem., 2008, 77, 51-76.

26.

Roy, R.; S. Hohng; T. Ha. A Practical Guide to Single-Molecule FRET, Nat. Methods,
2008, 5, 507-516.

27.

Zhao, R.; D. Rueda. RNA folding dynamics by single-molecule fluorescence resonanceenergy transfer, Methods, 2009, 49, 112-117.

28.

Förster, T. Zwischenmolekulare Energiewanderung und Fluoreszenz, 1948, 437, 55-75.

29.

Einstein, A. Strahlungs-Emission und -Absorption nach der Quantentheorie, Deutch.
Phys. Ges., 1916, 18, 318-323.

17

30.

Schuler, B.; E. A. Lipman; P. J. Steinbach; M. Kumke; W. A. Eaton. Polyproline and the"spectroscopic ruler" revisited with single-molecule fluorescence, Proc. Natl. Acad. Sci.
USA, 2005, 102, 2754-2759.

31.

Akiyama, B. M.; M. D. Stone. Assembly of Complex RNAs by Splinted Ligation,
Methods Enzymol., 2009, 469, 27-46.

32.

Fiore, J. L.; E. D. Holmstrom; D. J. Nesbitt. Entropic Origin of Mg2+-Facilitated RNA
Folding, Proc. Natl. Acad. Sci. USA, 2012, 109, 2902-2907.

33.

Fiore, J. L.; B. Kraemer; F. Koberling; R. Erdmann; D. J. Nesbitt. Enthalpy-Driven RNA
Folding: Single-Molecule Thermodynamics of Tetraloop–Receptor Tertiary Interaction,
Biochemistry, 2009, 48, 2550–2558.

34.

Holmstrom, E. D.; D. J. Nesbitt. Real-time infrared overtone laser control of temperaturein picoliter H2O samples: "Nanobathtubs" for single-molecule microscopy, J. Phys.
Chem. Lett., 2010, 1, 2264–2268.

18

Chapter 2

Experimental Methods

Each scientific discipline has its own set of tools that are frequently used to investigate
experimental problems. A single-molecule fluorescence microscopy tool kit is required for thevast majority of the scientific research described in this thesis. This chapter provides a detailed
description of most of the experimental techniques and fundamental scientific concepts that havebeen used to generate the contents of this work.
2.1

Experimental Observation
Nearly every scientific experiment requires some form of observable that can bemonitored to confirm or reject a particular hypothesis. A detailed and rigorous understanding of
this necessity is essential for an objective interpretation of the experimental outcome. As
mentioned previously, fluorescence is the preferred method of detection for most of the work
described in this thesis.
2.1.1

Molecular Fluorescence
Molecular fluorescence is a useful physical phenomenon in that it provides theexperimenter with the ability to very nearly observe chemical processes. Most generally,
fluorescence refers to the emission of light from a substance (e.g., molecule) that has absorbed
some form of electromagnetic radiation. This rather simple definition contains two profoundscientific processes—absorption and emission—that have been studied by many of the great
intellectuals of the past two centuries, perhaps most notably Albert Einstein(1). Absorption is aprocess that describes the interaction between light and matter, whereby the energy of an

19

absorbed photon (Ephoton = hc/λ) is transferred to the interacting matter (e.g., molecule). The
efficiency of this process is dictated by the physical properties of both the light, as well as, thematter and is characterized by a parameter called the absorption cross section (σ). This can bethought of as a nanoscopic area presented by the molecule to the photons, where the zerodimensional particles of light can only be described in terms of the average number of particles
per area. Absorbing a photon becomes more likely as the number of them occupying thismolecular cross section increases. The energy of the absorbed photon can be used to rearrange
the electronic configuration of the molecule to energetically excited state.
Radiative relaxation occurs when an excited state molecule returns to the ground state
and, in the process, emits lights. The energetic difference between these two states dictates theenergy content (i.e., color) of the emitted photon—higher energy photons are blue, low energy
photons are red. If this is the only available relaxation pathway to the ground state, then every
single absorbed photon will give rise to an emitted photon, resulting in a quantum efficiency (Φ)
of unity. However, there is another competing parallel pathway—non-radiative relaxation—that
allows excited state molecules to return to the ground state without emitting a photon. In thisprocess, the absorbed energy is released as heat, as opposed to a particle of light, which results inΦs of less than one. The Φ can also be defined by the rate constants associated with all of theexcited-state processes in the molecule (Eq. 2.1), where krad is the spontaneous rate constant forradiative relaxation (emission) and knrad,i is the rate constant for any of a number of non-radiative(i.e., non-emissive) relaxation processes.
Φ!=! k

krad
!
+
rad
i knrad,i

20(Eq. 2.1)

The rate constants can be used to characterize the excited-state fluorescence lifetime (τfluorescence,Eq. 2.2), which defines the average duration a molecule spends in the excited state (e.g., S1)
before returning to the ground state either though radiative or non-radiative relaxation.
τfluorescence = k

1
rad +

i knrad,i(Eq. 2.2)

These processes, and their associated time-scales, can be summarized in a Jablonski diagram(Figure 2.1).

Figure 2.1 Jablonski energy level diagram depicting absorption (excitation) and emission(fluorescence) of photons.(http://www.microscopyu.com/articles/fluorescence/fret/fretintro.html)
For most molecules in nature, non-radiative processes dominate their radiative
counterparts, giving rise to negligible Φs; it is only a very specific sub-set of molecules (i.e.,
fluorophores) with the right properties (i.e., large σ and high Φ) that have significant
fluorescence. Examples of organic fluorophores that are of direct relevance to this work are thecyanine (e.g., Cy3 and Cy5) and rhodamine (e.g., RhoB, and Rho6G) dyes. The relevant
photophysical properties (σ, τfluorescence, Φ, krad) of these molecules are summarized in Table 2.1.
Because the fundamentals of fluorescence are well understood, these fluorophores can be
attached, or coupled, to nucleic acids to probe specific characteristics of the labeled system. For21

example, one could attach a cyanine dye to a fictitious RNA of interest to determine where it islocated within a cell.

Table 2.1 Table of approximate photophysical properties associated with four fluorophorescommonly used throughout this thesis. All reported values are for free dye molecules in bufferedaqueous solutions, expect for Cy3-DNA*, where the conjugation to a nucleic acid significantly
increases the quantum yield and fluorescence lifetime of the fluorophore.
2.1.2

Forster Resonance Energy Transfer (FRET)
Indeed, fluorescence itself is a useful tool for locating the spatial position of a molecule.

However, in order to use this process as a read-out for questions more sophisticated than thoserelated to location, changes in fluorescence properties must be measured. Most frequently thesechanges are related to the above photophysical parameters (Table 2.1); of particular relevance tothis work are changes in the τfluorescence. The previous examples of non-radiative relaxation were
all intramolecular processes, however intermolecular processes can also significantly influencethe τfluorescence of a dye. One example of this is Forster (sometimes Fluorescence) Resonance
Energy Transfer (FRET). This phenomenon describes the non-radiative (i.e., non-emissive)transfer of energy from an energetically excited donor molecule to a neighboring acceptor
molecule via near-field dipole-dipole coupling. The rate constant for this energy transfer process
is dependent on a number of physical parameters (Eq. 2.3): (i) the relative orientation of the twointeracting dipoles (κ2), (ii) distance between the donor and acceptor fluorophores (r), (iii) the22

spontaneous rate constant for emission associated with the donor (krad,D), (iv) the spectral overlap
integral between the emission spectrum of the donor and the molar extinction spectrum of theacceptor (J), the index of refraction (n) of the medium containing the two fluorophores (typicallywater n = 1.33), and Avagadro’s Number (NA).
9k

ln 10 κ2 J

rad,D!!"#$ = 128
π5 n4 ! ! !

(Eq. 2.3)!

The orientation factor, κ2, is defined using Eq. 2.4,
κ2 =! cosθt -3cosθd cosθa2(Eq. 2.4)

where θt is the dihedral angle between the two planes associated with the transition dipole
moments of the donor emission and the acceptor absorption, and θd and θa are the angles
between the line-of-centers connecting the two fluorophores and the donor emission and acceptor
absorption transition dipole moments, respectively. This value can range from 0 to 4 depending
upon the relative orientation of the coupled dipoles(2). A value of 2/3 is often used, whichassumes that the time-averaged orientation of the two fluorophores isotropic. This assumption,
while generally quite sound, starts to break down for slowly rotating dipoles, like fluorescentproteins, and can be completely invalid for fixed, or oriented, dipoles.
The efficiency of energy transfer from the donor to the acceptor (EFRET) can be calculated(Eq. 2.5) in a similar fashion to the quantum efficiency (Eq. 2.1).
EFRET = kkFRETrad,D +kFRET + i knrad,D,i(Eq. 2.5)

In certain systems, the presence of an acceptor can significantly decrease (i.e., quench) theτfluorescence and Φ of the donor fluorescent dye. However, if the acceptor is also a fluorophore, theloss of donor fluorescence due to energy transfer is conveniently offset by the resulting
generation of excited-state acceptor fluorophores that can undergo typical spontaneous emission.

23

Again, a Jablonski diagram can be used to schematically illustrate the details of this process(Figure 2.2).

Figure 2.2 Jablonski energy level diagram depicting Fluorescence Resonance Energy Transfer(FRET). (http://www.microscopyu.com/articles/fluorescence/fret/fretintro.html)
The distance-dependence of FRET allows it to be used as a semi-quantitative ruler to measure
distances of approximately 10-100 Å. For the Cy3-Cy5 donor-acceptor FRET pair used
exclusively throughout this thesis, the distance corresponding to half-maximal EFRET (R0) is
approximately 53 Å; when the donor and acceptor are proximal (i.e., r < R0) EFRET is high, when
they are distal (i.e., r > R0) EFRET is low (Eq. 2.6).
R6

EFRET = 6 o 6
Ro +r(Eq. 2.6)

By attaching a FRET pair to a molecule of interest, not only can you determine its spatial
location, but you can also determine the distance between the two fluorophores, which provides
structural or conformational information about the molecule (Figure 2.3). Analogous to theexample mentioned above, a fictitious FRET-labeled RNA within a cell may, perhaps, reveal thatthe molecules are in a closed structure (high EFRET) in some areas of the cell and an open one(low EFRET) in other areas.
24

Figure 2.3 Distance dependence of FRET efficiency (EFRET) for a fictitious biological molecule.
Separation of the donor and acceptor fluorophores results in less efficient energy transfer.
2.2

Single-Molecule Confocal Microscope
Of course, the fluorescently labeled molecules are thousands of times smaller than thewidth of a human hair. A home-built single-molecule confocal fluorescence microscope is used
to facilitate observation of these tiny objects (Figure 2.4). The experimental system was firstbuilt and characterized by a previous postdoctoral researcher in the lab (Dr. Jose Hodak), withsome additional assistance from my former graduate student mentor (Dr. Julie Fiore). Details of
the experimental apparatus are contained in this section (2.2) of the thesis; the thesis of Dr. Julie
Fiore is also an excellent resource(3).
2.2.1

Excitation Path
The entire microscope system has been optimized for the use of the Cy3-Cy5 FRET pair,starting with the excitation source. A 532 nm (green) frequency doubled Nd:YAG laser is used toexcite donor (Cy3) fluorophores. This excitation source was chosen to facilitate spectral filteringof excitation light (532 nm) and Cy3 emission (λmax ≈ 570 nm). The excitation source is pulsed
at 20 MHz with a pulse width of ≈ 10 ps to enable the use of time-correlated single-photon
counting techniques (TCSPC, see section 2.2.4). A half wave plate (λ/2) and broadband

25

polarizing beam splitter are used to attenuate the laser power and ensure complete polarization of
the laser output. The 1 mm intensity diameter (1/e2) of the Gaussian laser beam is expanded to 12mm using a Keplerian telescope containing lenses with focal lengths of f1 = 25 mm and f2 = 300
mm, respectively. A former graduate student—Dr. Tom Baker—provides a great description ofproper techniques associated with measuring beam diameters in the experimental chapter of histhesis(4).

Figure 2.4 Cartoon cut-away diagram of the single-molecule microscope system used to conduct
the research associated with this thesis.
Each green laser pulse triggers a red laser pulse from a 635 nm (red) laser diode (< 100 ps
pulse FWHM) with an approximately 25 ns delay determined by the length of BNC cablebetween the two laser controllers. The time-delay and repetition rate of the two light sources
result in almost perfectly interleaved laser pulses. The incorporation of a red excitation source tothe experimental apparatus provides the ability to directly excite both donor and acceptor
fluorophores via Alternating Laser EXcitation (ALEX)(5-8), which ensures that biomolecules

26contain both donor and acceptor fluorophores. The red laser (635 nm) is used to efficiently excite
Cy5 (λmax ≈ 650 nm), while still being easy to spectrally filter from both Cy3 (λmax ≈ 570 nm)
and Cy5 (λmax ≈ 670 nm) fluorescence. The output of the red laser is directly coupled into asingle-mode fiber-optic cable to clean up the beam shape, as suggested by the manufacturer. The
light exits the cable via a beam collimator resulting in an intensity diameter (1/e2) of ≈ 10 mm.
The red and green laser beams are spatially overlapped using a dichroic mirror that transmits thegreen and reflects the red light. The two beams pass through a set of broadband neutral density
filters to attenuate the optical power before being directed into the back port of an Olympus IX70
inverted microscope base.
2.2.2

Microscope and Sample Stage
Once the red and green beams are inside the microscope, they are reflected upward

towards the back aperture (daperture = 9.25 mm) of the 1.2 N.A. water objective (UPLSAPO
60XW) via a custom 2-color dichroic mirror designed specifically to only reflect light from thetwo excitation sources (Di01-R532/635, Semrock, Inc). Because both the red and green beams
sufficiently overfill (i.e., intensity diameter (1/e2) > daperture) the back aperture, the opticalelements within the objective can effectively focus the light to a single diffraction-limited Airyfeature (Figure 2.5) at a working distance of 280 µm. The spatially dependent intensity of the
Airy feature can be expressed via Eq. 2.7, where J1 is a first-order Bessel function of the firstkind, r is the radius from the optical axis, λ is the wavelength of light, and N.A. is the numericalaperture the optical system.
I(x)
I0

=

2J1 !

2

x

, x≈

!!rN.A.$nλ(Eq. 2.7)

Accordingly, the radius that corresponds to the first zero of the Airy feature (rAiry) is
approximately equal to 0.61nλ/N.A., which corresponds to ≈ 360 nm for the present apparatus.

27

Figure 2.5 (left and middle) Visual depictions of the Airy disc associated with a diffractionlimited focus. (right) Graphical representation of the normalized radial intensity distribution (redsolid line, I(x)/Io) and the relative encircled power (blue dotted line, P(x)/Po). Note x = kasin(θ)=2πrN.A./λ, as used in Eq. 2.7. (http://en.wikipedia.org/wiki/Airy_disk).
The sample holder (Section 2.3.1) is supported directly above the objective by a 3-D
nano-positioning piezo-electric scanning stage, which is used to raster the sample in the plane of
the nanoscopic focus in order to generate a microscopic image of the sample (Figure 2.6).

Figure 2.6 Sample surface image of immobilized fluorescence molecules using the 3-D nanopositioning peizo-electric scanning stage (not shown).
This practice facilitates localization of fluorescent molecules. Because of the size and surface
density of the immobilized species, the fluorescent features (e.g., spots) associated with them are
diffraction limited and only sparsely populate the surface of the sample. The stage can be used toposition the sample such that a single fluorescent molecule will reside precisely at the center ofthe diffraction-limited focus. This exposes the fluorophores to the excitation sources resulting in
the absorption and subsequent emission of photons. The emitted fluorescence is collected by the28same objective (i.e., epifluorescence) and directed as a collimated beam of donor (Cy3) andacceptor (Cy5) fluorescence through the 2-color dichroic mirror, where it is focused by a tube
lens (f =80 mm) and directed out the side-port of the microscope by a mirror.
2.2.3

Emission path
The entire emission path, starting directly from the side-port of the microscope base, iscontained in a series of black matte optical tubes to prevent ambient room light from reaching thedetectors. A pinhole is placed ≈ 80 mm outside of the side port in the image plane of the tube
lens (f = 180 mm) within the microscope body. The aperture of the pinhole (dpinhole = 50 µm) isused to reject out-of-plane and off-axis. The 50 µm aperture is more than sufficient to capture thecenter spot of the Airy pattern after being magnified by the microscope objective (60 × dAiry ≈ 42µm). Notably, this central feature contains ≈ 83 % of the total power of the Airy pattern (Eq.
2.8).
P(x)
J0 !
=1
−
P0
x

2

−

J1 !
x

!

, x≈

!!rN.A.$nλ(Eq. 2.8)

On-axis fluorescence in the focal plane passes through the pinhole, where it is collectedby a lens and imaged onto the active area of the downstream detectors. Prior to reaching thedetectors, the fluorescence is spatially separated, first by polarization (horizontal and verticalwith respect to the laser table) using a polarizing beam cube (CVI PBSH-450-1300-100) andthen by color (e.g., donor and acceptor fluorescence) via dichroic mirrors designed to transmit
red and reflect green light (645DCXR, Chroma Technology). The last elements in the emission
path prior to the detectors are band pass and long pass filters on the donor and acceptor emission
paths, respectively. These optical elements facilitate rejection of photons from the excitation
sources (532 nm and 635 nm) and ensure that only Cy3 or Cy5 fluorescence reaches each of thefour detectors.

29

2.2.4TCSPC Detection
Four avalanche photodiodes (APDs, SPCM-ARQ-14, Perkins-Elmer Optoelectronics)

serve as the detection elements (1: donor horizontal, 2: acceptor horizontal, 3: donor vertical, and
4: acceptor vertical) for the Cy3 and Cy5 fluorescence. These remarkable devices provide theability to detect single-photons with high efficiency (≈ 65%) and enable time-correlated singlephoton counting (TCSPC) capabilities. A synchronization signal associated with the repetition
rate of the green laser (20 MHz) is directed to the TCSPC module (SPC-134, Becker & Hickl
GmbH) to track the laser pulses. Coincident with the detection of a single photon, the APD starts
a linear voltage ramp that stops when the next sync signal is received by the TCSPC module.
Because the ≈ 25 ns between laser pulses is much longer than the τfluorescenceof Cy3 and Cy5, theamplitude of the voltage and the known repetition rate of the green laser can be used to infer thearrival time (micro-time) of the photon relative to previous laser pulse that excited thefluorophore. The micro-times provide the ability to observe chemical dynamics on the 0.5 to 50
ns time scale (e.g., excited-state relaxation and rotational diffusion). Additionally, the TCSPC
module also keeps track of the arrival time of individual photons with respect to the start of theobservation period (macro-time). This provides the ability to observe the time-dependence of thephoton arrival rate for any arbitrary time-bin greater than 50 ns, making it possible to monitor
fluorescence changes resulting from longer time-scale chemical processes (e.g., translational
diffusion or biomolecular folding) ranging from hundreds of nanoseconds (10-7 s) to thousandsof seconds (103 s).
2.2.5

Temperature Control
The above single molecule apparatus enables observation of chemical processes via

fluorescence. The amount of information provided by this instrument is useful for tackling a30

wide range of scientific questions. However, precisely controlling the experimental sampletemperature can further increase this utility. Temperature is a useful experimental variable that,in conjunction with kinetics experiments, can be used to reveal the thermodynamic properties ofa system. To this end, the above experimental apparatus has been retrofit with two different
heating technologies, each capable of precisely and accurately controlling the sample
temperature. A previous graduate student (Julie Fiore) incorporated the more user-friendly stageheating capabilities, whereas the slightly more technical infrared-based optical heating
capabilities have been implemented as part of the research associated with this thesis.
2.2.5.1 Bulk Heating Methods
Instead of placing the sample holder (Section 2.3.1) directly on the 3-D nano-positioning
piezo-electric scanning stage, it is placed within an enclosed stage heater that, itself, rests on topof the scanning stage (Figure 2.7).

Figure 2.7 Diagram of the original, bulk, heating capabilities established by Dr. Julie Fiore(stage not shown).
Vacuum grease (Dow Corning) maintains efficient thermal contact between the sample holder
and the stage heater. The objective is almost completely thermally isolated from the stage andmust also be heated to minimize thermal gradients across the sample. This is accomplished by a31

resistively heated wire-based objective collar (Figure 2.7).Calibration of sample temperature
with respect to the set point of both the stage and objective heaters is necessary to ensure
accurate and precise temperature measurements. It is important to note that because of precious
optical elements in the objective, heating rates should not exceed 1 °C/min so as to minimize thepotentially damaging effects of thermal expansion. Furthermore, the temperature of the objective
should not exceed 50 °C according to the manufacturer. Because of the high heat capacity
associated with the objective, a 10 min wait is required for all of the elements to reach a steadystate temperature. In its current implementation, the bulk stage heating capabilities provide theability to reliably control the sample temperature (± 0.2 °C), though not without some
restriction—to use the words of my advisor, “There is no such thing as a free lunch!” Theseexperimental restrictions are primarily related to the following three concepts. First, biologicalsamples tend to rapidly degrade at elevated temperatures. This is particularly true for acid/baseand metal ion-catalyzed cleavage of the phosphodiester backbone in RNA(9,10). Althoughsample heating is desirable, heating the entire sample will result in many unwelcome side effects
related to sample degradation. Second, the large heat capacity of the objective mandates thatheating must occur on the time-scale of 10 min, which, unfortunately, makes taking datalaborious and time-consuming. Furthermore, these temporal constraints preclude the ability to
use rapid changes in temperature to induce particular conformational transitions in nucleic acids.
Lastly, thermal expansion of the objective limits the maximum attainable temperature, which
impedes the study of ultra-stable nucleic acid interactions that require high (> 50 °C)
experimental temperatures.

32

2.2.5.2 Infrared-based Heating
All of these hindrances associated with bulk stage heating can be alleviated thought theuse of optical heating methods. Specifically, focused infrared light resonant with a combination
band of the asymmetric and symmetric OH stretch in water can be used to vibrationally excite
water molecules (Figure 2.8). These excited molecules will rapidly deposit the excess energy
into the solution resulting in an increase in the sample temperature. A detailed description of theimplementation and characterization of these heating capabilities is the topic of Chapters 3 and 4,
which should be referenced for additional details.
In summary, 1443 nm continuous wave radiation is directly coupled into a single-mode
fiber at the output of the laser diode. The infrared light exits the fiber at the top of an optical cage
system via a beam collimator and propagates coaxially towards the sample and microscope
objective with a beam intensity diameter (1/e2) of ≈ 9 mm.

Figure 2.8 Cut-away cartoon of the infrared-based heating apparatus coupled to the existing
inverted confocal fluorescence microscope.
A lens (f = 100 mm), approximately 10 cm from the focal plane of the microscope objective, isused to focus the infrared light to an intensity diameter (1/e2) = 20 µm, which provides the33thermal energy for the infrared heating capabilities. The implementation of these methods
increase the range of accessible temperature to 20-90 °C, which represents a substantial
improvement from the 20- 50 °C range associated with bulk heating methods. The infrared
heating methods also result in substantially smaller heated volume (≲ 10-11 L) than conventional
heating methods (≳10-6 L), thus greatly reducing the effects of elevated temperatures on sample
degradation. Lastly, and perhaps most importantly, the optical modulation of temperatureprovides the ability to alter the sample temperature on a ms time-scale rather than the hundredsof seconds necessary with the previous implementations of temperature control on theexperimental apparatus.
2.3

Sample Preparation
Central to nearly every biological experiment is sample preparation, especially in systems

where inadequate sample preparation can lead to misleading experimental results. Although theexperiments described in this thesis are mostly insensitive to variations in sample preparation,some knowledge of this process is certainly useful. The following sections describe the processes
associated with preparing both the sample and the sample holders
2.3.1

Sample Holder
The single-molecule experiments described herein make use of different sample holder

variants that are all based on standard microscope cover slips. Corning No. 1½ 22 × 22 mm Zinc
Titania (ZT) cover slips (Catalog No. 2870-22) were used until they were discontinued in 2012.
Since then, Corning No. 1½ 22 × 22 mm Borosilicate (BS) cover slips (Catalog No. 2850-22)
have proven to be a less-than-desirable replacement due to fact they require more stringent
cleaning methods. Future efforts should be directed at finding a suitable replacement for theoriginal ZT coverslips. Historically, all cover slips were soaked in 70% nitric acid over night,

34

rinsed with HPLC grade-water, and dried with argon gas. This process is unnecessary for the ZT
coverslips, as many experimental observables are insensitive to the cleaning procedures.
However, this is not the case for the BS coverslips, which benefit from the nitric acid wash.
Given the University’s recent concern about both nitric acid and piranha cleaning methods (seeCU Boulder EHS), a simple methanol rinse has, thus far, proven to be sufficient for clean glass
surfaces.
The ready-to-use glass cover slips (either ZT or clean BS) can easily be incorporated into
one of three styles of sample holder (Figure 2.9): (i) the “open-air” sample holder (ii) “sandwich”
style, double-sided sticky tape sample holder, or (iii) PCTFE (polychlorotrifluoroethylene)
polymer sample holder. The majority of work conducted by a former graduate student (Dr. Julie
Fiore) made use of the PCTFE sample holders (see Dr. Julie Fiore’s thesis). Given the time andmaterial consumption required by these sample holders, both for cleaning and preparation, theother two, more crude and efficient samples holders quickly replaced them.

Figure 2.9 Image of the three types of sample holders used for fluorescence studies. See Section
2.3.1 for details.
The “open-air” style sample holder simply consists of a ready-to-use coverslip held in place by amachined aluminum coverslip holder. These were most often used for freely diffusing
experiments (Section 2.4.1) of free dye molecules or fluorescently labeled molecules in solution.
The “open-air” sample holders are also frequently used for alignment purposes. The “sandwich”
style sample holder consists of two pieces of Scotch Brand (3M) double-sided sticky tape, lying

35

parallel to each other, between a ready-to-use coverslip and a standard glass slide (1.0 mm thick
25×75 mm WVR microslides, No 48300-025). The open-ended ≈ 80 µm high channel, or tunnel,
resulting from this configuration, which typically runs perpendicular to the long axis of the glass
slide, provides the ability to flush, or more precisely, wick (via capillary action) different
solutions into and out of the sample holder. This “sandwich” style sample holder can besupported directly above the objective, coverslip-side down, by any two lightweight objects ofsimilar height (e.g., glass slides). Because of the ability to readily change solutions, these sample
holders were the desired choice for nearly all surface-immobilized and some freely diffusing
experiments involving fluorescent species.
2.3.2

Sample of Interest
Preparation of the actual experimental sample is obviously dependent on the type ofexperiment and, therefore, the style of sample holder. Freely diffusing experiments (Section
2.4.1) can be performed in the “sandwich” style sample holders, but are generally easier to
conduct using the “open-air” style sample holders. For experiments requiring freely diffusing
fluorophores, the dyes can be dissolved in HPLC grade H2O at dilute concentrations, typically
0.1 – 1000 nM, depending on the photophysical properties of the fluorophore. An approximately
40 µl drop of solution can be placed directly on the ready-to-use coverslips of an “open-air”
sample holder. These types of experiments are typically used for system alignment andcharacterization.
Unfortunately, a number of detrimental chemical transformations can occur when the
fluorophore is in the excited electronic state. The products of these excited state reactions (e.g.,
triplet states and conformational isomers) have been shown to be associated, either directly or
indirectly, with photophysical abnormalities. Molecular oxygen (O2) plays an important role in a36

number of these excited state chemical transformations. Of particular relevance is the strong
electrophile, single molecular oxygen (1O2), which rapidly reacts with the conjugated double
bonds of organic fluorophores, rendering them non-fluorescent. Singlet molecular oxygen has
also been known to oxidize nucleobases (e.g., guanosine), which can inhibit proper folding of
nucleic acids. For these reasons, two different photophysical stabilizing systems are used in orderto minimize the influence of O2. Specifically, a biological antioxidant/triplet state quencher—
TROLOX(11) (2 mM)—and an enzymatic oxygen scavenging system—PCA/PCD(12) (5 mM
and 60 nM, respectively)—are incorporated into all of the smFRET sample solutions. This
practice sufficiently increases the photostability of the fluorophores without influencing any ofthe measured properties associated with nucleic acid conformational transitions.
For experiments involving surface immobilized molecules (Section 2.4.2), the commonlyused chemistry of the biotin-streptavidin ligand-binding system, is employed to non-covalently
attach individual molecules to a ready-to-use coverslip at a surface density of less than 1
molecule per square micron. This is most efficiently accomplished using the “sandwich” stylesample holders. The sample holder is initially flushed with approximately 200 µl of 10 mg/mL
bovine serum albumin (BSA) containing 10% biotinylated-BSA and allowed to incubate for 10
min. This allows the BSA to non-specifically absorb to the surface of the glass coverslip. Then,
another approximately 200 µl of 200 µg/mL streptavidin is flushed through the sample holder
and allowed to incubate for 10 min. The incubation permits formation of the non-covalent ligand
binding interaction between biotin and streptavidin, which immobilizes streptavidin to thesurface of the coverslip. Conveniently, streptavidin is a tetrameric protein with four biotin
binding sites, which allows this protein to also bind biotinylated nucleic acids. Approximately
200 µl of a 100 pM solution of these biotinylated, fluorescently labeled molecules is flushed37

through the channel of the “sandwich” style sample holder to achieve the desired surface
densities (Figure 2.6, 2.10). After the nucleic acids have been immobilized to the surface of thecoverslip, the desired imaging conditions (including TROLOX and PCA/PCD) can be flushed
into the sample holder to permit prolonged observation of individual molecules.

Figure 2.10 Schematic representation (not-to-scale) of the commonly used biotin-streptavidin
surface immobilization technique. Approximately one out of ten BSA(blue ovals) molecules
contains a biotin (black triangle) moiety. Streptavidin (purple rectangle) has four binding sites
for biotin, which can be used to immobilize fluorescent nucleic acids to the glass coverslip.
2.4

Data Collection and Analysis
The single-molecule confocal fluorescence microscope, described above, is capable of anumber of advanced fluorescence spectroscopy techniques ranging from time-dependent
polarization anisotropy to fluorescence correlation spectroscopy. However, the majority of
research contained within this document is carried out using only two such techniques—freely
diffusing and surface immobilized experiments; both are well suited for single-molecule
experiments.
2.4.1

Single-Molecule Freely Diffusing
The single-molecule freely diffusing (sometimes referred to as “burst”) experiments

make use of a confocal microscope’s three-dimensional: (a) Excitation Probability Distribution(EPD)—determined by the microscope objective—and (b) Detection Probability Distribution

38

(DPD)—determined by the pinhole and detectors. The product of these two spatially dependent
probability distributions—called the molecule detection function (MDF)—is used to define a
region in space—termed the confocal volume—where the probability of detecting emitted
fluorescence is greater than 1/e2 (13). For the current microscope setup, this region isapproximately 0.36 femtoliters; one molecule in such a volume corresponds to a solution
concentration of approximately 5 nM. By working with concentrations of fluorescent species
much lower than this (i.e., 100 pM), one can ensure that most of the time the confocal volume isvoid of fluorescent species. However, when a fluorescent molecule stochastically diffuses
through this volume (Figure 2.11), fluorescence well above background will be observed.

Figure 2.11 Schematic for freely-diffusing single-molecule experiments.
The duration of the burst of fluorescence is obviously limited by the diffusion constant of thespecies in the solution; for the nucleic acids studied in this thesis, this amounts to approximately
1 ms in the confocal volume. The short residence times minimize the concern forphotobleaching, thus allowing for higher incident laser powers (i.e., 30-100 µW), which ensurethat the fluorescent bursts are well above background and contain a sufficient number of photons
for quantitative determination of EFRET. Conveniently for single-molecule FRET experiments,
the use of Alternating Laser EXcitation (ALEX) (see Section 2.2.1) provides the ability to39

determine the stoichiometry of the fluorescent busts such that only species with both a single
Cy3 and Cy5 are considered for data analysis(5-8,14).
Numerous 60-second time trajectories, each containing approximately 500-1000
fluorescents bursts can readily be acquired using in-house software—SPCImage.exe—written
originally by Dr. Vasiliy Fomenko using Lab windows/CVI(Figure 2.12). This software hassince been updated and modified by Dr. Thomas Baker, Dr. Julie Fiore, Dr. Larry Fiegland, andmyself.
Analysis of the data is performed using a second piece of in-house software also scripted
primarily by Dr. Vasiliy Fomenko—vffta64.exe (Figure 2.13). Binning the macro-time photon
arrival data at 1 ms (depending on the diffusion coefficient of the fluorescent species) allows for
single time-bin bursts of fluorescence. Count rate thresholds (20-35 kHz) can be assigned for
fluorescence bursts (Options > Burst Analysis) resulting from both the green and red lasers toensure that only Cy3-Cy5 labeled fluorescent species are considered for analysis.
Lastly, all of the dually labeled fluorescent bursts are compiled together in a histogram, which
depicts the distribution of EFRET values associated with the given experiment. The resulting shapeof the histogram, but not the amplitude, should be insensitive to the choice of red and green laser
ALEX threshold. The histograms can be analyzed a variety of ways to derive equilibrium data
and. More sophisticated techniques can even be used to extract kinetic data from such EFRET
histograms, as done by other members of the single-molecule community(15). The mostfrequently used method involves fitting the EFRET distributions to a sum of Gaussians with anadditional constraint determined by the total number of observed fluorescence bursts.

40

Figure 2.12 Images of the scanning (top) and time trajectory (bottom) windows of the
SPCImage.exe software used to acquire fluorescence data.
2.4.2

Single-Molecule Surface Immobilized
The second, but perhaps most powerful, operational mode afforded by the inverted

single-molecule scanning confocal fluorescence microscope involves acquisition of time-41

trajectories from individual surface-immobilized fluorescently-labeled nucleic acids (Figure2.10).

Figure 2.13 Image of the vffta64.exe software use to analyze the data acquired via the
SPCImage.exe program.
Because of the small area associated with the diffraction-limited focus of the excitation sources,
the sample must be rastered over the objective to generate an image of the surface immobilized
molecules (Figure 2.6). This can be accomplished using the SPCImage.exe software describedabove. Specifically, the software is used to control the 3-D nano-positioning piezo-electricscanning stage such that a 10 µm × 10 µm area of the sample is rastered in the plane of the focus
resulting in a 256 × 256 pixel image of the surface, where each pixel results from approximately
2 ms of fluorescence integration. The resulting RGB false-colored surface representation can beused to re-position the scanning stage such that a single fluorescent species resides directly in thefocus of the excitation sources. Using the SPCImage.exe software, time-traces can be acquired
from the individual fluorescent species for the specified duration of time (Figure 2.12). Analysisof these surface-immobilized trajectories is also accomplished using the vffta64.exe software

42

(Figure 2.13). Because of the time-tagged data storage method, the bin time can be adjusted,
post-acquisition, to achieve the desired balance of signal-to-noise and temporal resolution;
typical bin times range from 5-1000 ms depending on the experimental system. The software canalso be used to generate a corrected (16) EFRET trajectory which is then integrated over time togenerate a EFRET histogram for the specific time-trace. A dwell time threshold can be placed at an
EFRET value that approximately represents the minimum between the two distributions. The
software will generate a list of all of the threshold crossings and their corresponding dwell times.
The dwell times can then be sorted based upon the transitions that gave rise to them (i.e., high
EFRET → low EFRET, or low EFRET → high EFRET) and analyzed to determine the characteristic
time constant (or rate constant) that describes the process responsible for the threshold crossing
events.
2.5

Single Molecule Construct Design
Perhaps the single most important component of any single-molecule experiment is the

preparation of a fluorescently labeled construct. First and foremost, suitable locations forsynthetic modifications must be chosen so as to accurately report on conformational transitions
within the nucleic acid without significantly perturbing any of the structural or dynamical
properties of the molecule. This can be a particularly daunting task without the assistance ofbiochemical and structural data. Biochemical data can assist in identifying robust nucleotides
that can tolerate significant perturbations (e.g., mutations, insertions, and deletions), while still
maintaining wild-type activity. These locations are ideal for synthetic modifications likefluorescent dyes. Structural data can also be used to aid in construct design by providing asuitable model for intermolecular distances and the solvent-exposure of particular positions
within the nucleic acid. Ideally, the labeling sites should allow nearly free rotation of the43

fluorophores and the desired conformational transition should modulate the distance between the
fluorophores from >53 Å to <53 Å.
Once a construct has been conceptualized, synthesis must be carried out. For designs thatconsist of annealed oligonucleotides, the starting materials can be purchased from any of anumber of commercial sources capable of custom nucleic acid synthesis. However, often times,
more complicated steps are required to synthesize a suitable construct. For example, nucleic acidligation may be required to “join” together two (or more) oligonucleotides. There are a numberof useful (but challenging) ligation strategies that can be employed to accomplish such a task. Inhouse coupling of fluorophores to the nucleic acids may also be necessary for constructsynthesis. These techniques are often described and provided by the manufacturer of thefunctionalized fluorophores. Upon successful synthesis of an RNA construct, great care should
be taken to clean up and purify the product by removing undesirable fluorescent contaminants
and excess reactants. This is best accomplished using High Performance Liquid Chromatography(HPLC). Once purified, samples should be stored at -70 °C in 50 mM HEPES buffer with ~100mM monovalent cation concentration and 0.1 mM EDTA to prevent metal-ion catalyzed
cleavage of the nucleic acids. Aliquots from the original stock solution (µM) can be made at ~ 25
nM and stored at -20 °C for day-to-day experimentation. A more detailed description of theprocedures required to prepare each of the nucleic acid constructs covered in this thesis can befound in the experimental methods section of each of the corresponding chapters.

44

2.6

References

1.

Einstein, A. Strahlungs-Emission und -Absorption nach der Quantentheorie, Deutch.
Phys. Ges., 1916, 18, 318-323.

2.

van der Meer, B. W.; D. M. van der Meer; S. S. Vogel. Optimizing the Orientation Factor
Kappa-Squared for More Accurate FRET Measurements, In FRET – Förster Resonance
Energy Transfer, pp 63-104, Wiley-VCH Verlag GmbH & Co. KGaA.2013

3.

Fiore, J. L. (2012) Single-Molecule Kinetic and Thermodynamic Studies of CationFacilitated RNA Folding: Isolated Tertiary Interactions, In Chemical Physics, Universityof Colorado, Boulder.

4.

Baker, T. A. (2012) Confocal Microscopy Studies of Fluorescence Blinking of
Semiconductor Quantum Dots, Metal Nanoparticle Photogeneration, and Multiphoton
Photoemission from Thin Metal Films and Metal Nanoparticles, In Chemistry, Universityof Colorado, Boulder.

5.

Kapanidis, A. N.; T. A. Laurence; N. K. Lee; E. Margeat; X. Kong; S. Weiss.
Alternating-laser excitation of single molecules, Acc. Chem. Res., 2005, 38, 523-533.

6.

Kapanidis, A. N.; N. K. Lee; T. A. Laurence; S. Doose; E. Margeat; S. Weiss.
Fluorescence-aided molecule sorting: Analysis of structure and interactions byalternating-laser excitation of single molecules, Proc. Natl. Acad. Sci. USA, 2004, 101,
8936-8941.

7.

Lee, N. K.; A. N. Kapanidis; H. R. Koh; Y. Korlann; S. O. Ho; Y. Kim; N. Gassman; S.
K. Kim; S. Weiss. Three-color alternating-laser excitation of single molecules:
Monitoring multiple interactions and distances, Biophys. J., 2007, 92, 303–312.

8.

Lee, N. K.; A. N. Kapanidis; Y. Wang; X. Michalet; J. Mukhopadhyay; R. H. Ebright; S.
Weiss. Accurate FRET Measurements within Single Diffusing Biomolecules using
Alternating-Laser Excitation, Biophys. J., 2005, 88, 2939-2953.

9.

Forconi, M.; D. Herschlag. Metal ion-based RNA cleavage as a structural probe, Methods
Enzymol., 2009, 468, 91-106.

10.

Lonnberg, T.; M. Luomala. Intracomplex general acid/base catalyzed cleavage of RNA
phosphodiester bonds: the leaving group effect, Org. Biomol. Chem., 2012, 10, 67856791.

11.

Rasnik, I.; S. A. McKinney; T. Ha. Nonblinking and long-lasting single-molecule
fluorescence imaging, Nat. Methods, 2006, 3, 891-893.

12.

Aitken, C. E.; R. A. Marshall; J. D. Puglisi. An Oxygen Scavenging System for
Improvement of Dye Stability in Single-Molecule Fluorescence Experiments, Biophys. J.,
2008, 94, 1826-1835.

45

13.

Ruttinger, S.; V. Buschmann; B. Kramer; R. Erdmann; R. Macdonald; F. Koberling.
Comparison and accuracy of methods to determine the confocal volume for quantitative
fluorescence correlation spectroscopy, J. Microsc. Oxford, 2008, 232, 343-352.

14.

Joo, C.; H. Balci; Y. Ishitsuka; C. Buranachai; T. Ha. Advances in single-molecule
fluorescence methods for molecular biology, Annu. Rev. Biochem., 2008, 77, 51-76.

15.

Torella, J. P.; S. J. Holden; Y. Santoso; J. Hohlbein; A. N. Kapanidis. Identifying
molecular dynamics in single-molecule FRET experiments with burst variance analysis,
Biophys. J., 2011, 100, 1568-1577.

16.

Fiore, J. L.; J. H. Hodak; O. Piestert; C. D. Downey; D. J. Nesbitt. Monovalent andDivalent Promoted GAAA-Tetraloop–Receptor Tertiary Interactions from Freely
Diffusing Single-Molecule Studies, Biophys. J., 2008, 95, 3892-3905.

46

Chapter 3

Real-Time IR Overtone Laser Control of Temperature in Picoliter H2O Samples:“Nano-Bathtubs” for Single Molecule Microscopy*

3.1

Introduction
Temperature arguably represents the most critical experimental variable for detailed

thermodynamic/kinetic study of chemical and biological processes. Of particular interest,
temperature control at the focus of a confocal microscope objective offers significant potential
for kinetic exploration of nanoparticle(1,2) and biological(3-5) systems at the single molecule
level. Such temperature control can in principle be achieved from “bulk” methods by heating theentire sample, stage, and objective with an enclosed stage-heater and heated objective collar.
Though quite powerful, there are several important physical limitations to such a bulk heating
approach. First of all, the time constants for heating/cooling of the stage and objective can be
unacceptably long, requiring multiple minutes to achieve steady-state temperatures necessary forsingle-molecule fluorescence detection. Secondly, the speed of such bulk stage heating
approaches preclude use of powerful temperature jump methods to activate a specific process ofinterest, such as denaturation/renaturation of proteins, melting of nucleotide interactions,annealing of nanomaterials, etc. Furthermore, elevated objective temperatures (> 50 °C) cansoften optical cement and thereby destroy the precise lens alignment crucial to high numericalaperture photon collection. Finally, prolonged exposure to elevated temperatures may facilitate*

Adapted from: Holmstrom, Erik D.; Nesbitt, David J. J. Phys. Chem. Lett. 2010, 1, 2264-2268
47

unwanted degradation of the entire sample, which can decrease length of observation times andincrease background fluorescence.
Despite these challenges, there have been attempts to alleviate/overcome some of theshortcomings associated with bulk sample-heating, particularly with biological systems.
Resistive joule-heating in microfluidic systems has been used to increase sample temperatures to> 90 °C and thereby induce polymerase chain reaction; (6-8) however constraining the heating towithin a small sub-volume of the sample is difficult. Others have used the absorbance of water in
the near-IR region to directly heat the surrounding medium(9-15). The use of IR light provides
optical control of the heated volume, as well as large, programmable temperature changes overshort periods of time. As a result, IR laser overtone heating is well poised to remedy a number ofshortcomings associated with traditional stage-heating methods. Specifically, the use of resonant
near-IR laser excitation of H2O permits optically controlled, uniform heating of picoliter
volumes in a well-defined confocal geometry. Furthermore, any residual transmitted IR laser
intensity can be selectively blocked with high specificity by attenuation with a thin bulkabsorbing H2O filter. As a result, the combination of uniform sample heating and exponential
rejection of the heating light permits temperature-dependent studies of kinetics/thermodynamics
while maintaining continuous, high fluorescence sensitivity necessary for single-molecule
studies.
The thrust of the present work describes an IR-absorption based heating approach
coupled with single-molecule fluorescence resonance energy transfer (smFRET) microscopy tosimultaneously heat and observe folding of single RNA molecules. To accomplish thermal
control of RNA samples, IR light is used to vibrationally excite the first overtone band (vOH =2←0 at 1445 nm) of the OH-stretch in water (Figure 3.1). This non-contact heating method48

makes it possible to systematically control the temperature of small volumes of aqueous solution.
The result is a confocal fluorescence microscopy apparatus capable of broad temperature control(T = 20-90 °C), high accuracy (± 0.5 °C) and heating rates in excess of 104 °C/sec, which offers
enormous potential for thermodynamic/kinetic studies of conformational dynamics of
biomolecules at the single-molecule level. The reliability and accuracy of this technique aretested by using the IR-absorption based heating to reproduce thermodynamic folding studies of
the GAAA tetraloop−receptor from the Tetrahymena ribozyme(16).

Figure 3.1 Cutaway diagram of IR absorption based heating apparatus. Inset shows theabsorption coefficient (α) of water in the vOH = 2!0 overtone band region near 1.45 mm.
3.2

Results and Discussion
To achieve IR-absorption based temperature control of aqueous solutions, a fiber-coupled

1445 nm continuous-wave laser has been incorporated into an inverted fluorescence microscope(Figure 3.1) with a 532 nm pulsed laser excitation source. This apparatus is capable of color,polarization, and time-resolved single-photon(16). In the studies described herein, an XYZtranslation stage controlled optical-cage is used to focus (f = 100 mm) IR light down to a 17 µm(full width half maximum) diameter spot in the focal plane of (and coaxial with) the microscope49

objective. The IR laser power (PIR) is controllable between 0-160 mW, with typical peak incident
laser intensities of Io » 47 kW/cm2. At l = 1.445 µm, the absorption coefficient, α(l), is equal to= 30.3 cm-1 (17). For an H2O sample depth of ≈ 150 mm, this corresponds to a Beers Law
absorbance of A ≈ 1.05 (base e) and nearly uniform local heating throughout the ≈ 35 pL sample
volume. For an aqueous sample dominated by the heat capacity of water (Cp(H2O) = 4.184
J/K/cm3), this model predicts a maximum heating rate ([ ∂T/∂t](t = 0) = [α(l) × I° ]/Cp) of 3.4 x 105°C/sec.
A microscope objective collects the fluorescence from a donor/acceptor fluorophorelabeled RNA construct. The fluorescent photons are separated from the excitation source anddirected towards the detection system using a dichroic mirror. Before detection, the fluorescentphotons are spatially separated by color and polarity using various optical elements toaccommodate FRET experiments (see Fiore et al.(16)). Detection of fluorescent photons isaccomplished with single-photon avalanche photodiodes and a time-correlated single-photon
counting (TCSPC) module(16). A thin 3 mm water filter located under the objective is used toprevent excess IR light not absorbed by the sample from reaching the detection system. Infrared
light is attenuated with essentially complete efficiency (T ≈ 8 × 10-10), and yet with minimal lossof sample fluorescence (T ≈ 0.9).
An important associated challenge is accurate measurement of the sample temperature,
which is accomplished using TCSPC methods to measure the temperature-dependent
fluorescence lifetime (τfluor) of rhodamine B (18). To establish reliable calibration over the fullrange of interest (20-90 °C), the rhodamine B lifetimes are measured (i) in situ with TCSPC
methods and (ii) benchmarked against studies in a temperature-controlled fluorometer. The
fluorescence decay curves in both regimes are least squares fit to a convolution of single-

50

exponential decay and a well-defined instrument response function to determine τfluor as afunction of temperature.
Previous experiments exploiting this property of rhodamine B have focused mainly oneither τfluor or fluorescence intensity, with the resulting empirical calibration curves typicallyinterpolated as a means of inferring the solution temperature(13,19-21). We can take thisanalysis one step further by considering a realistic Arrhenius kinetic model for the temperaturedependence of rhodamine B. Specifically, it has been shown that the quantum yield (QY) of
rhodamine B approaches unity as the temperature decreases(18). Additionally, it is also known
that the rate constant for the non-radiative relaxation from the excited state (knrad) is strongly
temperature-dependent and vanishes at low temperature(18,22). Based on the relationshipbetween decay rate constants and fluorescence quantum yield (QY = krad/[krad + knrad]), theseobservations suggest the temperature dependence of τfluor to arise from the non-radiative rate, i.e.
knrad(T). This can be simply modeled as an activated process, with the temperature dependence ofknrad(T) dictated by the well known Arrhenius equation, i.e., knrad(T) = Aexp(-Ea/kT).
Analysis and results from such an Arrhenius model are summarized in a standard Arrhenius plotof ln[knrad(T)] versus 1/T (Figure 3.2). Over the range of temperatures observed, ln[knrad(T)]decreases linearly with 1/T, which when fit to the Arrhenius equation yields an activation energy(Ea) for accessing the non-radiative decay channel of 7.2(1) kcal/mol. This is qualitatively
consistent with previous Arrhenius analyses of non-radiative decay in methanol (Ea = 5.2(4)kcal/mol), ethanol (Ea = 5.0(4) kcal/mol) and 1-propanol (Ea = 4.1(5) kcal/mol), which is clearly
indicative of a reduction of Ea with decreasing solvent polarity (23). Of particular relevance tothe present work, this expression provides physically-motivated calibration of τfluor as a function51

of solution temperature, and thereby a non-invasive measure of temperature for picoliter volumesof H2O.

Figure 3.2 Arrhenius analysis of the non-radiative component knrad(T) for temperature-dependent
rhodamine B fluorescence rates. By explicitly measuring kfl = krad + knrad (T) and modeling knrad(T) as a thermally activated Arrhenius process, one can use TCSPC and the Arrhenius equation
as a calibration curve for temperature.
As a quantitative demonstration of such a IR-absorption based heating method, we havealso investigated equilibrium thermodynamics for RNA folding at the single-molecule level (16).
Specifically, a donor/acceptor labeled RNA construct and smFRET are used to study thetemperature-dependent folding/unfolding dynamics of the GAAA tetraloop-receptor tertiary
interaction. Most importantly, this permits a direct comparison with previous single-molecule
measurements of enthalpy and entropy based on bulk microscope stage-heating.
The equilibrium constant as well as the individual rate constants for folding/unfolding aredetermined using previously established methods(24). Figure 3.3 depicts a van’t Hoff plots of R
ln[Keq] versus 1/T obtained from both the present study (IR-absorption heating) as well as
previous work (bulk stage heating). Results from this work (ΔHº = 18(2) kcal/mol, ΔSº = 58(6)52cal/mol/K) and bulk stage heating experiments (ΔHº = 17(2) kcal/mol, ΔSº = 56(5) cal/mol/K)
are clearly in excellent agreement, with all thermodynamic quantities indistinguishable within
experimental uncertainty. This represents strong confirmation that the present IR-laser
absorption based heating methods can be used to obtain high quality kinetic/thermodynamic dataon single-molecule samples.

Figure 3.3 Van’t Hoff plot of R ln[Keq] vs 1/T for single molecule FRET studies of RNA folding
provides ΔHº and ΔSº for the GAAA tetraloop−receptor tertiary interaction. Note the excellent
agreement between the laser-overtone and stage-based heating techniques.
One critically important advantage of the IR laser heating method is the high temporalcontrol of sample temperature, which we can address more quantitatively from the data. To
determine a maximum rate and upper limit time-scale for heating, fluorescence from rhodamine
B is measured as a function of time during abrupt heating/cooling cycles (Figure 3.4). To obtainsufficient accuracy in the fluorescence lifetime information, τfluor at each time point isdetermined from a least-squares exponential decay fit to 2000 photons, which at maximum
fluorescence rates corresponds to a minimum bin time of ≈ 20 ms. The prompt decrease in τfluor
reflects a correspondingly rapid increase in knrad(T), which from the Arrhenius expression
53

translates into an IR laser-induced jump of ΔT = 23.2(5) ºC. Interestingly, the characteristic rate
for laser heating above ambient temperature is currently limited by the minimum bin time, withthe data clearly establishing an upper limit for the heating process (theat < 20 ms). It is also worth
noting that the ability to monitor confocal fluorescence continuously throughout the temperature
jump event highlights a critical kinetic advantage of the present narrow band IR laser absorption
heating technique over broad band IR-induced heating methods(12).

Figure 3.4 Time scale for IR-absorption based heating. By monitoring rhodamine B fluorescence
lifetimes, one can directly measure characteristic times associated with laser heating of thesample. The characteristic heating time is currently limited by the time binning resolution of thedata (< 20 ms), which is slower than, but consistent with, predictions (see Section 3.2 fordetails).
This analysis can be explored further based simply on conservation of energy and
Newton’s Law of cooling. If we at first assume no heat flow into/out of the illuminated sample
volume, conservation of energy relates the time-dependent deposition of thermal energy (∂(ΔQ)/∂t = α(l) × Io) to the time-dependent change in local temperature (∂(ΔT)/∂t = Cp(H2O) ∂(ΔQ)/∂t).
In this approximation, the temperature derivative is given by ∂(ΔT)/∂t = (α(l) × Io )/Cp(H2O),
which for 50 mW of IR light focused to an area of A = π(17 mm)2/4 at 1445 nm (α(l) = 30.3 cm54

1

) predicts a maximum initial rate of ≈ 1.1 × 105 °C/sec. It is worth noting that because of therelatively small absorbance per unit length, and therefore uniform heating conditions, thisprediction is rather insensitive to the sample depth.
Of course, this initial rate is not maintained indefinitely due to thermal conduction andloss of heat to the surrounding medium, resulting in a steady-state temperature increase. Thistemperature change can be analyzed via Newton’s law of cooling, ∂(DT)/∂t ∝ –ΔT, which canbe integrated to yield a single-exponential rise/fall to an asymptotic steady-state temperature, i.e.,

ΔT(t) = ΔT∞ × [1 – exp(-t/ τheat)]. This time constant (τheat) will be a function of thermalconduction efficiency away from the laser heated volume. However, t heat can be simply reexpressed from the expression above to yield τheat = (ΔT∞)/[∂(ΔT)/∂t](t = 0) in terms of (i) the
maximum initial rate of temperature change ([∂(ΔT)/∂t](t = 0)) and (ii) the asymptotic temperaturerise (ΔT∞), which can be experimentally measured for a given PIR. Note that since both ΔT∞ and[∂(ΔT)/∂t](t = 0) are approximately proportional to intensity, τheat will be largely insensitive to theincident PIR.
More quantitatively, for a 43 mW IR laser focused to a sample area of A = 235 mm2 inwater, the asymptotic temperature rise is empirically found to be ΔT∞ = 23.2(5) °C. From theabove expression, this predicts τheat ≈ 0.25 ms, which is substantially faster but neverthelessconsistent with the experimentally determined upper limit. We are currently extending thesestudies to digitally switched laser outputs, faster fluorescence rates, and shorter bin times toconstrain this limit further. However, we anticipate that some of this discrepancy may be a result
thermal gradients and heat flow. By way of evidence, the heating and cooling time constantsfrom such a first-order model are predicted to be equal, whereas experimentally the cooling time
is observed to be considerably longer (τcool ≈ 320(20) ms) and easily measurable with a 20 ms

55

bin time. One interpretation is that on the time scale of heating (> 20 ms) the heated volume
closely resembles the profile of the incident IR laser beam. However, over longer times heat candiffuse radially away from the pseudo-column of water giving rise to a broad volume of heatedwater. As such, this large volume of water will cool down more slowly than it takes the small
volume of water to heat up. Unfortunately, this broad heated region will expose more surface
tethered molecules to elevated temperatures and potentially contribute to unwanted sample
degradation and background fluorescence, but is still significant improvement from bulk sample
heating. Nevertheless, this simple first-order analysis of rapid temperature changes suggests
intriguing prospects for temperature jump capabilities and single-molecule kinetics on the msec
time scale. Specifically, this technique coupled with smFRET provides the ability to observe, in
real-time, intermediate states in the melting pathway of large non-coding RNAs.
3.3

Conclusions
In summary, we have presented results from a confocal method with high spatial (30 µm)

temporal (< 20 ms) and thermal (± 0.5 K) control of aqueous smFRET samples based on
excitation of the OH-stretch overtone with 1445 nm laser light. The benefits of this non-contact
method of resonant IR laser heating are (i) much larger dynamic temperature range compared tomore traditional stage-heating methods, as well as (ii) the ability for continuous sensitive
monitoring of fluorescence at the single molecule level throughout the heating event. Thesetemperatures increases can be accurately calibrated from the temperature-dependent fluorescence
lifetimes of rhodamine B, making it particularly useful for quantitative studies of temperaturedependent kinetics/thermodynamics in a variety of systems.
Indeed, such resonant IR-laser absorption based heating techniques have very recently
been adapted to wide-field microscopy experiment (10,11) demonstrating the applicability of the56technique to single-cell in vivo studies. This work has furthered the application of IR absorption
based heating to the field of single-molecule microscopy. In particular, the prospects for
temporal control of picoliter sample temperatures down to the msec time scale with resonant
absorption of the residual IR flux offers access to temperature jump experiments performed at
the single molecule level and monitored in real-time.
3.4

Experimental Methods
The three-piece donor/acceptor labeled GAAA tetraloop−receptor construct, as well as all

solution preparations, annealing and tethering protocols are as described previously;(16) only
experimental modifications necessary for the current study are presented herein. Samples holders
are created with standard glass slides and coverslips. Double-sided tape is placed across a clean
glass slide creating two closely spaced parallel channels and sealed with a clean glass coverslip.
In the first channel, fluorescence measurements of 500 nM rhodamine B (in water) underconstant power illumination are used to determine knrad(T) with < 1% error, from which thesample temperature can be calculated from the Arrhenius expression calibration curve (Figure3.2). FRET trajectories from a series of single RNAs immobilized in the second channel are thencollected under identical constant power illumination, simply by translating the glass slide
assembly. As a control, independent tests with rhodamine B in both channels reveal notemperature difference within experimental uncertainty (± 0.5 K) that could potentially arise
from variations in the glass or sample-holder preparation. For all of the above experiments, themicroscope stage is not temperature controlled and thus is confined to the ambient room
temperature. As such, all temperature changes are a result of increases in laser power.

57

3.5

References

1.

Li, S.; K. Zhang; J. M. Yang; L. W. Lin; H. Yang. Single quantum dots as local
temperature markers, Nano Lett, 2007, 7, 3102–3105.

2.

Valerini, D.; A. Creti; M. Lomascolo; L. Manna; R. Cingolani; M. Anni. Temperaturedependence of the photoluminescence properties of colloidal CdSe/ZnS core/shell
quantum dots embedded in a polystyrene matrix, Phys. Rev. B, 2005, 71, 235409.

3.

Bartley, L. E.; X. W. Zhuang; R. Das; S. Chu; D. Herschlag. Exploration of theTransition State for Tertiary Structure Formation Between an RNA Helix and a Large
Structured RNA, J. Mol. Biol., 2003, 328, 1011–1026.

4.

Hohng, S.; T. J. Wilson; E. Tan; R. M. Clegg; D. M. J. Lilley; T. J. Ha. Conformational
flexibility of four-way junctions in RNA, J. Mol. Biol., 2004, 336, 69–79.

5.

Kato, H.; T. Nishizaka; T. Iga; K. Kinosita, Jr.; S. Ishiwata. Imaging of thermal activationof actomyosin motors, Proc. Natl. Acad. Sci. USA, 1999, 96, 9602–9606.

6.

de Mello, A. J.; M. Habgood; N. L. Lancaster; T. Welton; R. C. R. Wootton. Precise
temperature control in microfluidic devices using Joule heating of ionic liquids, Lab on a
Chip, 2004, 4, 417-419.

7.

Hu, G. Q.; Q. Xiang; R. Fu; B. Xu; R. Venditti; D. Q. Li. Electrokinetically controlled
real-time polymerase chain reaction in microchannel using Joule heating effect, Anal.
Chim. Acta, 2006, 557, 146–151.

8.

Zhao, Q. L.; Q. A. Huang; Y. C. Lin. Design and numerical analysis of a joule-heatinginduced continuous-flow polymerase chain reaction microchip, J. Micro-Nanolithogr.
MEMs and MOEMs, 2009, 8, 021101.

9.

Braun, D.; A. Libchaber. Trapping of DNA by thermophoretic depletion and convection,
Phys. Rev. Lett., 2002, 89, 188103.

10.

Ebbinghaus, S.; A. Dhar; D. McDonald; M. Gruebele. Protein folding stability anddynamics imaged in a living cell, Nat. Methods, 2010, 7, 319-323.

11.

Schoen, I.; H. Krammer; D. Braun. Hybridization kinetics is different inside cells, Proc.
Natl. Acad. Sci. USA., 2009, 106, 21649-21654.

12.

Zhao, R.; E. Alemán; D. Rueda. Laser Assisted Single-molecule Refolding, Biophys. J.,
2009, 96, Biophys. Soc. proceedings 9a.

13.

Slyadnev, M. N.; Y. Tanaka; M. Tokeshi; T. Kitamori. Photothermal temperature control
of a chemical reaction on a microchip using an infrared diode laser, Anal. Chem., 2001,
73, 4037–4044.

58

14.

Mao, H. B.; J. R. Arias-Gonzalez; S. B. Smith; I. Tinoco; C. Bustamante. Temperature
control methods in a laser tweezers system, Biophys. J., 2005, 89, 1308–1316.

15.

Peterman, E. J. G.; F. Gittes; C. F. Schmidt. Laser-induced heating in optical traps,
Biophys. J., 2003, 84, 1308–1316.

16.

Fiore, J. L.; B. Kraemer; F. Koberling; R. Erdmann; D. J. Nesbitt. Enthalpy-Driven RNA
Folding: Single-Molecule Thermodynamics of Tetraloop–Receptor Tertiary Interaction,
Biochemistry, 2009, 48, 2550–2558.

17.

Wieliczka, D. M.; S. S. Weng; M. R. Querry. Wedge shaped cell for highly absorbent
liquids: Infrared optical constants of water, Appl. Opt., 1989, 28, 1714–1719.

18.

Karstens, T.; K. Kobs. Rhodamine B and rhodamine 101 as reference substances forfluorescence quantum yield measurements, J. Phys. Chem., 1980, 84, 1871–1872.

19.

Muller, C. B.; K. Weiss; A. Loman; J. Enderlein; W. Richtering. Remote temperature
measurements in femto-liter volumes using dual-focus-Fluorescence Correlation
Spectroscopy, Lab on a Chip, 2009, 9, 1248-1253.

20.

Robinson, T.; Y. Schaerli; R. Wootton; F. Hollfelder; C. Dunsby; G. Baldwin; M. Neil;
P. French; A. Demello. Removal of background signals from fluorescence thermometry
measurements in PDMS microchannels using fluorescence lifetime imaging, Lab on a
Chip, 2009, 9, 3437–3441.

21.

Ross, D.; M. Gaitan; L. E. Locascio. Temperature measurement in microfluidic systems
using a temperature-dependent fluorescent dye, Anal. Chem., 2001, 73, 4117–4123.

22.

Arbeloa, F. L.; T. L. Arbeloa; M. J. T. Estevez; I. L. Arbeloa. Photophysics of
Rhodamines - Molecular-Structure and Solvent Effects, J. Phys. Chem., 1991, 95, 2203–
2208.

23.

Casey, K. G.; E. L. Quitevis. Effect of solvent polarity on nonradiative processes inxanthene dyes: Rhodamine B in normal alcohols, J. Phys. Chem., 1988, 92, 6590.

24.

Downey, C. D.; J. L. Fiore; C. D. Stoddard; J. H. Hodak; D. J. Nesbitt; A. Pardi. Metal
ion dependence, thermodynamics, and kinetics for intramolecular docking of a GAAA
tetraloop and receptor connected by a flexible linker, Biochemistry, 2006, 45, 3664–3673.

59Chapter 4

Pulsed Infrared-Heating Studies of Single Molecule DNA Duplex Dissociation Kinetics andThermodynamics*

4.1

Introduction
Over the years, single-molecule fluorescence techniques have proven to be quite

powerful by allowing researchers to monitor the kinetics and dynamics of individual
biomolecules in real-time. These techniques have resulted in a number of important scientificadvances regarding a wide variety of biological systems(1). One of the most common
applications of single-molecule fluorescence, Förster Resonance Energy Transfer (FRET), hasbeen particularly advantageous for the observation of distinct conformational transitions(2,3).
Although direct observation of biomolecular dynamics provides valuable kinetic and mechanistic
insight into the particular conformational transitions, even more information can be obtained by
combining such single-molecule FRET (smFRET) experiments with control of otherbiophysically relevant variables such as applied force and temperature(4-11). For example, theaddition of explicit temperature control to smFRET experiments can provide thermodynamic
information not only on the free energy change, but also the enthalpy and entropy changes
associated with the observed processes(4,9). In particular, efficient integration of high resolution
spatial and temporal control of temperature with existing single-molecule fluorescence
technology represents a significant advance and offers novel quantitative thermodynamic details
about biomolecular dynamics (12).
*Adapted from: Holmstrom, Erik D.; Dupuis, Nicholas F.; Nesbitt, David J. Biophys J. 2014,
106, 220-231
60

The most widely used method with which to control the temperature of an aqueous
sample on a single-molecule fluorescence microscope is though the use of an objective-stage
heating system(4,7,9). However, there are three major problems associated with such a heating
technique: (i) a limited range of accessible temperatures (<45 °C) resulting from temperature
restrictions on high numerical aperture objectives, (ii) large heated volume determined by thesize of the sample, and (iii) long equilibration times limited by thermal conduction. Recently, anumber of techniques have been developed that use focused near-infrared light to directly(10,1216) or indirectly(17,18) modulate the temperature of aqueous samples. The indirect heating
methods are based on infrared absorptions by thin metal films on glass substrates, which thentransfer heat to the aqueous solutions. The direct heating techniques all rely on the fact that watermolecules have a modest ability to absorb near-infrared light(19), which excites various
vibrational modes of the liquid water and effectively increases the local temperature. Of specificinterest in this work is the absorption band near 1440 nm corresponding to the (ν1ν2ν3) = (1,0,1)
combination state for one quantum of both symmetric and asymmetric OH stretching excitation.
These infrared-heating techniques can alleviate many of the limitations associated with theobjective-stage heating systems, specifically by providing a broader range of available
temperatures and vastly superior spatio-temporal control of temperature(10,12,18,20). Thoughindirect infrared heating techniques have been well-described(17), none of the direct techniques
have been quantitatively characterized, which represents the first major aim of this work.
As a second target, we focus on specific applications in single molecule biophysics.
Although the emergence of infrared-based heating methods has alleviated many deficiencies of
conventional stage heating techniques, it also has revealed experimental restrictions associatedwith continuous observation of single-molecules at elevated temperatures (e.g., > 45 °C). Two

61

particularly important restrictions arise from: (i) the temperature-dependent dissociation of the
non-covalent bond between biotin and streptavidin (21) and (ii) the temperature-dependent
quantum yields of common single-molecule organic fluorophores(22,23). Due to the unusuallyhigh binding affinity of biotin-streptavidin (Kd ≈ 5×10-14 mol/L) and the extremely slow
dissociation rate (kdiss ≈ 3.8×10-6 s-1 at 37 °C), this ligand binding interaction has been widelyused in single-molecule experiments for surface immobilization to microscope coverslips(24-27).
However, as is necessarily the case with all activated rate processes, the rate constant for
dissociation of the biotin-streptavidin interaction increases exponentially with temperature,
which can therefore result in rapid depletion of surface immobilized molecules. Key advances inthis area of specialized surface immobilization techniques have been fueled by powerful `click’
chemistry, which can be used to covalently attach single molecules to sample substrates insituations that require stable surface immobilization (28,29).
However, there is one universal difficulty associated with performing smFRET studies atelevated temperatures that represents a more serious experimental problem. In particular, allconventional organic dyes have fluorescence quantum yields that are dramatically reduced athigh temperatures. This is predominantly due to rapid growth in the non-radiative decay rate
constant, which competes efficiently with the radiative pathway and therefore makes observationof single fluorophores challenging at temperatures > 45 °C. There have been a variety of recent
efforts that help address this problem, ranging from: (i) the use of semiconductor nanoparticles(e.g., quantum dots) with much higher absorption cross sections, (ii) more efficient collection offluorophore emission by immobilizing individual molecules on colloidal spheres (30), and (iii)
development of intrinsically brighter fluorophores by conjugation with photophysical stabilizing
agents(31). Although such techniques represent possible directions for addressing issues of62

fluorophore quenching at elevated temperatures, they also add significantly to experimental
complexity, particularly for single molecule studies of biophysical interest. Despite theseadvances, there remains a considerable need for alternative microscopy methods that can offer
access to biomolecular dynamics at temperatures where visualization of common organic
fluorophores is challenging. The second aim of this paper is to describe ongoing efforts to help
address this issue.
This article is organized into two major sections corresponding to the two parallel aims
described above. The first thrust describes a detailed characterization of an all-optical infraredlaser based heating technique (12) for single molecule microscopy, which offers quantitative
analysis of the thermal, spatial, and temporal characteristics. When compared with conventional
microscope stage heating methods, three advantages regarding optical temperature control areapparent: (i) larger range of accessible temperatures, (ii) extreme localization of thermal load topicoliter volumes and (iii) orders of magnitude faster heating rates. As a complementary second
thrust, the high spatial/temporal control of the infrared heating is exploited for high-temperature
smFRET kinetic measurements. In particular, we present results from a “proof-of-principle”
demonstration, whereby rapid heating/cooling cycles are used to obtain detailed
kinetic/thermodynamic information on the dissociation of 21 bp DNA duplex. Interestingly, theresults clearly reveal non-monoexponential temporal behavior in the dissociation kinetics that areused to develop a simple kinetic model based on simultaneous “fraying” from each end of the
DNA duplex. Additionally, such a model is capable of describing duplex dissociation for both
long (>10 bp) and short (< 10 bp) DNA sequences.

63

4.2

Overtone Infrared Laser Based Heating Technique
Details of the infrared heating system used throughout the present study can be found inpreviously published material(12) and can be quickly summarized here. Specifically, the TEM00
Gaussian beam profile from a continuous wave laser diode with peak wavelength of 1443 nm isfiber-coupled into an optical cage system containing a beam collimator (output spot size, ω =4.5(3) mm) and focusing lens (ƒ = 100 mm) to form a suitably small waist (ωo = 10.0(4) mm,
FWHM ≈ 17 µm) for local heating near the focal plane. A precision three-axis translation stage
then translates the optical cage with respect to the diffraction-limited focus of a 532 nm
excitation laser beam (Figure 4.1a), which illuminates the sample in the opposite direction
through the objective of an inverted fluorescence microscope.

Figure 4.1 Schematic representation of the infrared heating apparatus illustrating the coaxiallyaligned foci of the infrared (red) and excitation (green) lasers within a standard sample holder(not to scale). The intensity plot depicts the experimentally measured radial intensity profile atthe focus of the infrared laser (ωo = 10.04(4) µm).

64

Beers law absorption (α1443= 30.3 cm-1) of infrared radiation induces vibrational OH stretch
combination band excitation (ν1, ν2, ν3) = (1,0,1) of the solvent water molecules, which relax on
the sub-ns time scale and thus yield a rapid increase in solution temperature in the laser focal
region. The following experiments have been performed with such an apparatus toexperimentally quantify each of the three major proclaimed advantages associated with infrared
heating: (i) larger range of temperatures, (ii) smaller heated volumes, and (iii) faster heating.
4.2.1

In Situ Measurement and Dynamic Range of Accessible Temperatures
One crucial aspect of the infrared heating apparatus is clearly the magnitude of theinduced temperature change. As described previously (12), these temperatures can be accurately
obtained by measuring the temperature-dependent fluorescent lifetime [τfluor(T)] of Rhodamine
B(32,33) when the heating and excitation foci are spatially overlapped. The essence of themethod relies on the fact that kfluor(T) is simply the sum of a temperature-independent radiative
process, krad, and a temperature-dependent non-radiative relaxation, knrad(T)(34). Furthermore,
knrad(T) can be accurately modeled using Eyring transition state theory (Figure 4.2, inset),reflecting that non-radiative relaxation is enhanced by tilting the peripheral tertiary amino groups
away from the plane of conjugation (34). If we invert this procedure, experimental extraction ofknrad(T) for Rhodamine B provides an in situ “thermometer” for the local temperature in thesample region.
Specifically, time-correlated single-photon counting (TCSPC) methods are used toacquire the fluorescence decay curve of Rhodamine B in the confocal detection volume of the
532 nm excitation laser. The kfluor(T) is then determined by single-exponential fits convoluted
over the instrument response function of the excitation source, with knrad(T) obtained bysubtracting the known krad value from kfluor(T). The resulting knrad(T) can be used to predict

65confocal temperatures to within ≈ 0.5 °C, based on the previously determined Eyring transition
state parameters for Rhodamine B in water (Figure 4.2, inset)(12). Over the range of incident
infrared laser powers (Plaser = 0-135 mW) focused to a waist of w0 = 10.0(4) µm at a distance 15µm above the glass cover slip, the present experimental apparatus allows for rapid,
systematically controlled temperature increases from ΔT = 0 - 80 °C, as illustrated in Figure
4.3a. These results unambiguously demonstrate a range of experimental temperatures
significantly larger than accessible by conventional stage-objective heating systems.

Figure 4.2 Fluorescent decay profile for Rhodamine B as a function of temperature. The
linearity of the fluorescence decay over three orders of magnitude attests to the singleexponential behavior of Rhodamine B. The inset represents the Eyring analysis of thetemperature-dependent non-radiative rate constant for Rhodamine B (see Section 4.2.1 fordetails), which is used to determine the temperature rise in the diffraction-limited confocal
volume of the sample.

66

4.2.2High Resolution Spatial Control of Temperature
The second major advantage associated with infrared heating methods is the high degreeof spatial localization for the induced temperature changes. In order to quantify this, we firstcharacterize the intensity distribution of the IR-heating laser at the beam waist. This can beaccomplished by measuring the IR laser light transmitted though a 5 µm circular pinholetranslated on a piezoelectric stage. The resulting plot of measured intensity vs. lateraldisplacement (Figure 4.1a) clearly depicts a TEM00 beam, which can be nicely fitted with aGaussian beam waist of ωo = 10.04(4) µm. This is also in very good agreement with expectationsof ωo = 10.3 µm, based on the propagation of a Gaussian beam at the fiber collimator output (ωo= 4.5(3) µm) that is focused to a waist by a 100 mm focal length lens.
To experimentally determine the steady-state radial temperature-profile, the optical cage
system, and thus the IR laser light, is translated horizontally away from the coaxial position
established by the excitation beam. Rhodamine B fluorescence lifetimes are then monitored by
TCSPC as a function of distance on the translation stage micrometer. The measured values ofkfluor(T) are then used to calculate ΔT as a function of the radial separation between the heatingand excitation sources, as summarized in Figure 4.3b. The data clearly demonstrate that understeady state heating conditions, the temperature increase is localized in the radial dimension(FWHM ≈ 55 µm). Additionally, the data also establish that the radial temperature profile issignificantly broader than the incident laser beam. At distances near (i.e. < 5 µm) the axis ofheating, the temperature gradient (≈ 0.1 °C/µm) is much smaller than the experimental
uncertainty associated with temperature determination (≈ 0.5 °C), which conveniently results inan essentially uniform temperature profile near the heating axis. It is important to note that thetemperature begins to gradually decreases at distances further from the heating axis. This slow

67

radial decay of temperature is consistent with the inverse logarithmic dependence of the radial
temperature profile associated with a one-dimensional temperature source and is a result of theelongated column of heat provided by attenuation of the focus infrared beam as it propagates
through the sample.

Figure 4.3 Experimentally measured steady state temperature rise (ΔT) as a function of (a)incident laser power as well as (b) radial and (c) axial distances from the laser focal point. The
solid red region in (b) represents the radial intensity profile of the infrared laser beam scaled tothe maximum ΔT. Note the clear broadening of the temperature profile beyond the laser
excitation region, which arises largely from thermal diffusion away from the heating axis.
By way of further characterization of the spatial temperature profile, ΔT has beenmeasured along the vertical axis of the infrared laser, as summarized in Figure 4.3c. It isinteresting to note that the measured temperature profiles are relatively flat with a gradual68maximum near the midpoint, and therefore only a small vertical gradient exists between the two
glass boundary surfaces. The reason for this is that overtone absorption strengths for water lie in
an ideal intermediate regime, whereby the 85 µm sample thickness is appreciably smaller than
the 1/e optical penetration depth (330 µm), and yet not so small as to non-linearly attenuate laser
propagation through the sample. As a result, the on-axis thermal heat deposition is nearly linearwith distance, which implies nearly uniform heat load in each axial volume element. Most
importantly, it is worth noting that the spatial characterization of heated volume demonstrates
that the present apparatus confines the temperature rise to extremely small volumes (≈ 10-10 L).
This represents a significant improvement relative to conventional bulk heating methods andfurthermore, with sufficiently dilute sample conditions, extendable down to the heating of asingle molecule construct.
4.2.3

Rapid Temperature Modulation
The last major advantage offered by infrared overtone laser heating is control of sample

temperature in the time domain, which is only limited by diffusion of heat. To provide
expectations for the magnitude(s) of such heating and cooling times, we consider the following
elementary physical model. The rate of deposition of thermal energy into a laser illuminatedwater cylinder of height Dl and cross sectional area A and bounded by an ambient temperature
heat bath (T0) is∂Q/∂t = αΔl Ilaser,(Eq. 4.1a)

where α is the normal base e absorption coefficient. By conservation of energy, this thermal
deposition must equal the time-dependent change in temperature (∂(T – T0)/∂t = ∂ΔT/∂t) divided
by the heat capacity (Cp AΔl) of the sample volume V = AΔl. This immediately yields the simple
equation69

(∂ΔT/∂t)0 = αIlaser/[Cp A](Eq. 4.1b)for the instantaneous rate of heating at t = 0. Furthermore, Newton’s Law of Cooling implies that
this initial rate of temperature increase (°C/s) must be linearly proportional to the asymptotic
temperature differential (ΔT∞) when steady state conditions are achieved, with coefficient ofproportionality given by the exponential rise time. The combination of these relations predicts acharacteristic 1/e time for cooling (or heating) given byτheat = ΔT∞ /(∂ΔT/∂t)0,(Eq. 4.1c)

One simple qualitative prediction from Eq. 4.1c is that since both (ΔT/∂t)0 and ΔT scale∞

approximately linearly with laser power, the time constant for heating (or cooling) of a localized
sample surrounded by material at T0 should be insensitive to incident laser power and depend
only on the sample heat capacity undergoing this ΔT∞. A second more quantitative prediction isthat for typical conditions of laser power (Ilaser ≈ 135 mW), absorption cross section (α = 30.3cm-1), and beam area (A ≈ ω02 ≈ 10-6 cm2), the initial heating rates for the sample volume areremarkably swift, on the order of 1 x 106 °/s! Finally, for a typical asymptotic temperature gain
under such conditions of ΔT = 80 °C, Eq. 4.1c suggests a 1/e time constant for heating on thesubmillisecond time scale.
In order to determine the experimental time-scales associated with heating and coolingaqueous solutions using focused infrared light, time-dependent trajectories of freely diffusing
Rhodamine B (575 nM) have been collected as a function of heating duration (Δt). For a given
Δt, each trajectory exhibits a rapid rise to steady-state temperature ΔT , at which point the∞deposition of thermal energy into system is perfectly balanced by loss of heat to the surroundings(e.g., objective, stage, air). At the end of the heating duration, the heat source is turned off andthe sample rapidly cools back to T0 to complete a single heating/cooling cycle (Figure 4.4a).

70Since the data are collected by TCSPC, the trajectories can be sorted into bins of 100 consecutive
photons, with each bin converted into a exponential fluorescence decay rate kfluor(T) by
maximum-likelihood estimation (35). As describe in Section 4.2.1, these kfluor(T) values can becorrected for the purely radiative contribution and converted into temperatures trajectories ΔT(t)from the knrad(T) Arrhenius expression for Rhodamine B (Figure 4.2). The heating/coolingbehavior of such a temperature trajectory is shown in Figure 4.4a for a heating duration of 500
ms and Plaser = 60 mW, which clearly reveals an extremely sharp increase in the sample
temperature.

Figure 4.4 Real time observation of the infrared heating and cooling behavior, as obtained bytime-correlated single photon fluorescence measurements of Rhodamine B in the confocal
excitation region. (a) Sample temperature trajectory (black ⊗’s) resulting from a 50 mW heating
pulse with a duration of 500 ms. The solid lines represent multi-exponential fits used to calculate
the half-maximal times for heating (solid red) and cooling (solid blue). (b) A linear-log plot
demonstrating the dependence of cooling time constant on the heating duration, which suggeststhat the system achieves a steady state thermal distribution on the few second time scale.

71

Though too rapid to resolve with 100 photon time bin steps, the time to half-maximal
temperature increase (τ½heat) suggests a heating time constant ≤ 1 ms, in good qualitative
agreement with the above predictions.
Closer inspection of Figure 4.4a reveals that the cooling profile after the laser is switched
off is noticeably broadened with respect to the rise. Specifically, for a laser heating duration of
Δt = 500 ms, τ½cool ≈ 22(2) ms whereas τ½heat ≈ 1 ms. It is worth stressing that this is notinconsistent with the equal times predicted above from Newton’s Law of Cooling (Eq. 4.1c). The
reason is clearly evident in Figure 4.3b. Lateral thermal conduction away from the laser
illumination region during the approach to steady state necessarily increases the surrounding
temperature to well above T0. It is then the presence of this additional “heat reservoir” adjacentto the laser heated region that slows the eventual diffusive cooling process back to ambient
values. As expected from such a bulk diffusion process, these decays also deviate considerablyfrom single exponential, which makes characterization by τ½ values a more useful concept.
One clear prediction from this model is that the cooling times should be strongly dependent on
the heating duration. Such a compilation of data is presented in Figure 4.4b, which displays thecooling time ( τ½cool ) vs. duration of the heating laser (Δt). The results (in log time units) areconsistent with an approximately proportional growth in τ½cool with Δt, that eventually saturate atsome steady state value for the cooling process. The intercept of this initial linear region is 3(1)ms, which is qualitatively consistent with Newton’s Law of Cooling expectations that τ½heat ≈τ½cool for sufficiently short heating durations. Most importantly, such exceptionally short valuesfor τ½heat (and, at sufficiently small Δt, also for τ½cool) represent an enormous improvement (>104
fold) from the typical time scales of minutes associated with conventional stage-objective
heating methods. As will become clear in the following section, this offers several critical72

advantages for single molecule studies of biophysical kinetics, in particular enabling hightemperature single-molecule kinetic measurements without the need for temperature-stable
fluorescent dyes.
4.3

A Test Application: Single Molecule Kinetics of DNA Duplex Dissociation
As clearly outlined in the introductory sections, the use of IR overtone laser based

methods for controlled heating of picoliter volumes within an aqueous samples has considerable
potential for applications in a confocal single molecule microscopy. In particular, the above
discussion has revealed at least two fundamental complications associated with continuous
observation of individual, fluorescently labeled molecules at elevated temperatures: (i) thetemperature-dependent rate of dissociation for the biotin-streptavidin surface immobilization
technique (21,36) and (ii) the temperature dependent fluorescent lifetime of commonly usedsingle-molecule fluorophores (22,23). Because of the novel ability to rapidly modulate the local
temperature of aqueous solutions, pulsed heating experiments can in fact be used to circumvent
both of these difficulties. On the one hand, rapid T modulation achieves a enormously reduced
duty cycle for surface heating over bulk heating techniques, which dramatically reduces parallel
loss of the single molecule biotin–strepavidin attachment. On the other hand, the rapid heating
and cooling also greatly limits kinetic effects due to time spent in transition, which permits
subsequent FRET dye interrogation under cool, high quantum yield conditions. The following
pulsed heating experiments, based on a similar proof-of-principle demonstration for laser
assisted single-molecule refolding (LASR)(10), are used to demonstrate the ability to make hightemperature kinetics measurements of DNA duplex dissociation, tethered to a coverslip withbiotin-streptavidin interactions, without the need for thermally stable fluorophores.

73

4.3.1

Laser Heating Induced Duplex DNA Dissociation Kinetics
In order to exploit these IR laser heating methods to investigate the temperature-dependent dissociation kinetics of duplex DNA, a synthetically modified DNA construct has
been designed with two highly stable conformations—duplex or closed—that can be easily
distinguished via FRET (Figure 4.5a).

Figure 4.5 (a) Diagram of the duplex DNA fluorescent construct used for the dissociation
experiments. Green and red stars represent the Cy3 and Cy5 FRET pair used in the experiments.
The solid cyan line corresponds to the PEG18 internal linker within the construct to easily allow
the construct to form the closed conformation. (b) ALEX surface image (10 µm × 10 µm) prior
to and (c) after exposure to a pulse of infrared light. White circles represent the mask used tomeasure survival probability (see Section 4.3.1 for details).

74

The duplex is formed by hybridization of two oligonucleotides—reporter and splint. The reporter
oligo has: (i) a 6 base pair self complementary sequence separated by a (PEG)18 linker, (ii) Cy3
and Cy5 fluorescent dyes and (iii) a 3’ biotin to permit tethering to the surface via biotinstreptavidin interactions. The splint oligo simply contains 21 nt that are complementary to thereporter oligo. Formation of the duplex conformation is accomplished by hybridization of thereporter and splint oligos, which spatially separates the Cy3 and Cy5 dyes, resulting in low
FRET efficiency (EFRET ≈ 0.1, green). Upon dissociation of the splint, the reporter oligo is able torapidly form the closed conformation, whereby the first six nucleotides at the 5' end quickly basepair with nucleotides 7-12 at the 3' end of the PEG linker (Figure 4.5a). The transition fromduplex to closed conformations greatly reduces the radial distance between the Cy3-Cy5 FRET
pair, resulting in more efficient FRET (EFRET ≈ 1.0, red). The basic experimental technique istherefore to use a raster-scanning confocal microscope to image the FRET states under room
temperature conditions, before and after the IR laser has induced transient heating of the sample
up to a tunable but well characterized temperature for a given time duration.
Prior to a IR laser induced heating episode, alternating laser excitation methods (ALEX,
see Materials and Methods) are used to scan images (10 µm × 10 µm) of the sample surface,which reveal that nearly all of the dually labeled surface-immobilized molecules are in the (lowFRET state (green) duplex conformation (Figure 4.5b). Upon exposure to the IR laser heating
pulse (Δt =10 s, T=72.3(5) °C) centered on the image, the same surface shows that nearly every
duplex has undergone dissociation, as indicated by the abundance (> 90%) of molecules in theclosed (red, high EFRET) conformation (Figure 4.5c). As mentioned previously, the nearly
uniform temperature profile near the heating axis conveniently results in all of the molecules in
the 10 µm × 10 µm images experiencing the same temperature. Additional evidence for the75

spatially localized heating and duplex dissociation is demonstrated more dramatically in FRET
images of larger 80 µm × 80 µm scans (as shown in Figure 4.6) where the temperature gradientis no longer negligible. These larger images reveal a clear dividing line between the inner
circular region with dissociated duplexes (red, high FRET) and the surrounding area with fully
intact duplexes (green, low FRET). The reason is that duplex constructs in the inner circularregion have been heated to temperatures with sufficiently large rate constants to have significant
dissociation probability during the 10 s heating duration.

Figure 4.6 ALEX scanned image (80 µm × 80 µm) of the fluorescent DNA duplex constructs
after heating the center of the image to 72.3(5) °C for 10 sec. Note the clear circular region ofhigh FRET molecules (red), corresponding to duplex constructs that have undergone
dissociation, while the surrounding low FRET species (green) remain in the duplex
conformation.
As a result, most molecules heated by the infrared laser reside in the closed conformation (red,high FRET), while those around the periphery remain in the duplex conformation (green, low
EFRET). Long-term exposure to elevated temperatures is capable of inducing duplex dissociation
in a 21 bp DNA double helix, with the same duplexes more or less indefinitely stable at room
temperature. Therefore, we exploit the excellent spatial, temporal and thermal control of the IR
overtone heating laser to systematically tune the (i) temperature change and (ii) heating duration76

(Δt) in order to extract the temperature-dependence of the DNA duplex dissociation rate
constant.
The degree of duplex dissociation for a given heating duration (Δt) and steady-state
temperature (T) can be quantified via the single molecule survival probability, S(Δt,T), corrected
for loss of construct from view due to other channels such as photobleaching and disruption of
the biotin-streptavidin linkage. To extract S(Δt,T), one generates a mask (Figure 4.5b) that
identifies the number of individually immobilized molecules prior to the heating pulse with both
Cy3/Cy5 dyes and in the duplex conformation (Nduplex[0]). After exposure to the heating cycle,
the sample area is overlaid with the same mask (Figure 4.5c), in order to determine the numberof remaining molecules in the duplex conformation (Nduplex[Δt,T]) as well as the very small
number N’ (where N’ << Nduplex[0]) removed via parallel pathways such as photobleaching or
irreversible dissociation of the biotin –strepavidin linkage. Corrected for these loss channels(e.g., N’duplex [0] = Nduplex[0] – N’), the survival probability S(Δt,T) can simply be calculated via.
S(Δt,T) = Nduplex[Δt,T]/ N’duplex[0](Eq. 4.2)

The survival probability is monitored as a function of heating duration (Dt) and temperature (T)in order to probe the underlying kinetics mechanism of temperature-induced dissociation of a
21bp DNA duplex.
In order to correctly analyze these heating pulse experiments, one must make two kinetic
assumptions. The first is that the duplex dissociation kinetics are negligible at room temperature,and only contribute during the heating episode. This is readily confirmed by the fact that surface
immobilized constructs do not change FRET states over many hours in the absence of theheating laser. The second assumption is that the dissociation event is irreversible, with theliberated splint strand unable to re-form the duplex conformation. Since duplex association is a77

bimolecular process, this assumption is clearly valid, as the dissociated splint strand quickly
diffuses into the bulk to form a solution of vanishingly low concentration.
Pulsed-heating experiments are performed to measure the survival probability as afunction of the heating pulse steady-state temperature (T = 55.2(5) to 64.9(5) °C) and duration(Δt = 50-1000 ms). The results of these experiments clearly demonstrate that measured values of
the S(Δt,T) remain near unity for sufficiently short heating durations (< 100 ms) regardless ofpulse temperature. As the heating duration increases, S(Δt,T) decreases with a distinctly
sigmoidal shape, i.e., with the initial ∂2S/ ∂2t < 0 and a clear inflection point uncharacteristic ofpurely single exponential decay kinetics. The impact of heating duration on the kinetics is quite
clear from Figure 4.7a, with the time scale for achieving half-maximal survival probabilities
decreasing rapidly with increasing temperature. It is worth stressing that such a heating protocol
makes it possible to obtain kinetic information at elevated temperatures using standard
fluorescent dyes, since the fluorescence photons are harvested only under room temperature
conditions where high quantum yields have been reestablished.
We next consider the simple kinetic models necessary to describe these observed S(Δt,T)
plots for each of the four heating series. First of all, it is important to note that for any
dissociation process over a single rate-limiting barrier (e.g. duplex → hairpin). For such a
process, the survival probability would be described by a simple single-exponential decay (Eq.
4.3) with a rate constant, kdiss(T), that depends on the steady-state temperature of the heating
pulse.
S(Δt,T ) = e−Δt kdiss (T )(Eq. 4.3)

However, this model would predict a conventional convex shape (i.e., with initial ∂2S/∂2t > 0)lacking an inflection point and inconsistent with the data. Instead, the experimental survival78

probabilities clearly reveal an anomalous “lag time” at short heating durations (Figure 4.7a),which necessarily requires description by a higher than first order kinetic model. The simplest
such model is that of consecutive first-order reactions (e.g., D0 → D1 → C) where D0 and D1
represents some partially dissociated duplex conformation (i.e., still low FRET), whereas C
corresponds to the spectroscopically distinguishable closed (high FRET) conformation accessible
only by dissociation of the full duplex.

Figure 4.7 (a) DNA duplex survival probability data, S(Δt,T), as a function of heating duration,
Δt, at four steady-state heating pulse temperatures. Dashed lines represent least squares fit to aconsecutive first order reaction model, which yields n = 4.1(3) sequential reaction steps with anassumed identical rate constant. Notice the rapid increase in this rate constant with increasing
temperature, which implies a large endothermic component the transition state barrier. (b) Eyringtransition state analysis of the temperature-dependent rate constant, suggesting a highly
endothermic (ΔH‡ = 50(10) kcal/mol) and entropically rewarding (ΔS‡ = 100(20 cal/mol/K)
component of the free energy barrier for each of the fraying events.

79

In such a kinetic model, the “lag time” arises from the first order kinetics of D0 needing to form
D1 prior to forming the detected species C. Even more likely, there could be a series of n suchintermediate species (e.g., D0 → D1 ⋅⋅⋅→ Dn → C), which impact both the lag time as well as thecooperative sharpness of the sigmoidal behavior. If all n of the rate constants associated with thesteady-state temperature of the heating pulse, k(T), are all nearly identical, the solution to this nstep first order consecutive kinetic model can be solved exactly to yield

S(Δt,T ) =Γ(n,k(T )Δt)Γ(n)(Eq. 4.4)

where Γ(n) and Γ(n,k(T)Δt) are the gamma function and incomplete gamma function,
respectively. Successful examples of the application of sequential chemical reaction models tobiological systems include both DNA helicases(37,38), as well as strand exchange induced by
RecA(39). Application of this model to the present experiments involves fitting the data in
Figure 4.7a to extract the two fitted parameters: (i) the rate constant for each of the steady-state
temperatures associated with the infrared heating pulse, k(T), and (ii) the number of consecutive
steps, n. The results of a least-squares analysis of the data yield n = 4.1(3), which in the context
of such a model would suggest that there may be four consecutive steps prior to complete duplex
dissociation. Furthermore, the values of k(T) increase dramatically with relatively small increasein temperature, which would be consistent with the four sequential steps having to surmountstrongly endothermic free energy barriers. Other more complex kinetic models can of course fit
the data. However, the success of this simple consecutive first order reaction picture with n ≈ 4
proves a useful starting point, from which insights into the duplex dissociation event may be
developed further below.

80

4.4

Duplex Dissociation Model
Since the discovery of the dsDNA structure (40), there has considerable effort directed

toward understanding the fundamental kinetics and thermodynamics for duplex dissociation andassociation events (41). A significant amount of work has been directed at characterizing theforward association process, in particular highlighting the importance of pre-helical structures foroligonucleotide strands forming the duplex(42-44). For corresponding studies of the duplex
dissociation process, however, there has been much less consensus regarding the explicit
dissociation mechanism, with conflicting reports of single-exponential(45-47), stretchedexponential(48), and bi-exponential kinetic behavior(49,50) for different oligo length systems
and experimental conditions. The present single molecule heating results confirm that the DNA
duplex dissociation kinetics are distinctly non-single exponential for a 21 bp DNA duplex.
Specifically, the “lag time” dissociation behavior observed in these studies is fundamentallyinconsistent with that of an isolated rate process over a single transition state barrier. Instead, thedissociation kinetics are more correctly described by an first order consecutive model with n ≈ 4matching rate constants that increase rapidly with temperature.
This suggests a simple, physically motivated mechanism for duplex dissociation;
specifically, “fraying” of the 21 bp DNA duplex can occur from either of the two ends of thehelix. Before developing this model further, it is first important to note the pitch of the DNA
double helix is about 10 bp per 360° (51) and largely restricted by base-stacking interactions(52).
Thus, for a DNA duplex longer than 10 bp, simple structural constraints make it physicallydifficult to dissociate without major disruption of the helical pitch. This alone would suggest that
duplexes longer than 9 bp must initially peel apart (i.e., “fray”) at one or both ends to generate asufficiently short duplex that can then undergo rapid dissociation in a single step. For the 21 bp

81

DNA duplexes in this study, this would require fraying of at least 12 bp. Although fraying canoccur entirely from one end, dissociation preceded by symmetrical fraying of at least 6 base pairs
from each end would be much more highly favored by simple Poisson considerations.
Specifically, the statistical ratio between N base pairs fraying from two ends vs 2N base pairs
fraying from one end is readily shown to be (2N)!/(N!)2, which for N = 6 is already 924:1.
Similar ideas in the literature have been used to help interpret dissociation(45) and endfraying(53-55) dynamics of DNA duplexes. These studies of end-fraying dynamics attemperatures well below the melting point of the DNA duplex have been able to measure smallchanges in the terminal base pairs that occur on the 10-7 s time-scale. Of course, the present
analysis associated with the n-step consecutive first-order kinetic model is clearly not sensitive toindividual bp fraying steps, but instead appears to be consistent with approximately four fraying
steps with 3-5 base pair disruptions in each (see Figure 4.8). For the four consecutive fraying
events, Poisson considerations suggest that fraying from both ends would be six-fold times more
likely than fraying entirely from one end. Additionally, with more than 3 base pair disruptions ineach of the observed fraying events, it is not surprising that the time-scale for this process isorders of magnitude slower than what has been measured for single base pair fraying
dynamics(53,54). Therefore, it is reasonable to suggest that the 21 bp DNA duplex used in thesepulsed heating experiments dissociates via the following pathway: Two fraying events, of 3-5 bp
each, must occur at the two ends of the DNA duplex, giving rise to a highly frayed intermediatewith only a small number (1-9) of remaining base pairing interactions. This intermediate is thenable to dissociate in a rapid (i.e. non-rate-limiting) step into the two individual oligonucleotides,
as schematically depicted in Figure 4.8.

82

We note that this highly simplified analysis of the duplex dissociation kinetics is based ona series of rate-limiting steps due to fraying from both ends of the helix to fewer than 10
remaining bp, followed by rapid dissociation of the remaining duplex. Recently published singlemolecule experiments have investigated the association/dissociation kinetics of smaller DNA
duplexes (6-9 bp) as a function of temperature (56).

Figure 4.8 Model depicting the “consecutive fraying at both ends” mechanism for duplex
dissociation. The colored bars represent short segments of the 21 bp DNA duplex that are either
associated (green) or dissociated (red). The transparent black numbers correspond to the relative
abundance of each partially dissociated species as predicted by a binomial distribution. After
four consecutive fraying events, the single remaining associated (green) segment of the duplex issufficiently frayed to undergo a final rapid dissociation step (see Section 4.4 for details).
The results of these experiments strongly support the assumption of rapid single-exponential
survival probabilities for short DNA duplex that at elevated temperatures. Specifically, if one

83extrapolates to the higher temperatures (65 °C) in the present study, the temperature-dependent
single-exponential rate constants for duplex dissociation would be > 104 s-1. This is alreadyconsiderably faster than the range of rate constants obtained in Figure 4.7 and thus consistent
with the notion of rate-limiting symmetrical fraying to the center from both ends, prior to a rapid(non-rate limiting) dissociation step. As a final topic, the measured temperature-dependence ofthe n ≈ 4 sequential fraying rate constants can be used to extract thermodynamic information
about the fraying process itself. Specifically, from Eyring transition state theory, theexperimentally measured duplex dissociation rate constant is given by‡‡‡k(T ) = υ e (-ΔG kT ) = υ e (-ΔH kT ) e (-ΔS k )(Eq. 4.5)

where υ is some characteristic vibrational attempt frequency for the fraying event. We cantherefore formulate an Eyring transition state plot of ln[k/υ] vs 1/T to provide key additional
information on the enthalpy (slope) and entropy (intercept) components of the free energy
barrier. Such an analysis is shown in Figure 4.7b, which predicts a large enthalpic transition statebarrier of ΔH‡ = 50(10) kcal/mol and is consistent with breaking several base pair interactions ineach of the four fraying events for the DNA duplex. Indeed, for a mean dissociation enthalpy of≈ 8.4 kcal/mol/bp obtained from nearest neighbor modeling(57), this would be consistent with anaverage of ≈ 6(1) base pair disruptions in each of the 4 consecutive fraying processes. Similarly,
the corresponding entropy change from such an Eyring plot is found to be large and positive, ΔS‡= 100(20) cal/mol/K, which from a nearest neighbor modeling of the average entropy (22.4
cal/mol/K/bp) would suggest disruption of ≈ 4.5(9) base pairs in each of the 4 fraying events.
Although such an analysis is clearly not intended to be quantitative, these values are at leastconsistent with end fraying of the DNA duplex down to a small of base pairs prior to a final(non-rate-limiting) dissociation event. Clearly an interesting next step would be to further

84

explore such dissociation kinetics as a function of sequence and length of the DNA duplexes,
which would be necessary to further refine such a multi-step consecutive fraying mechanism forduplex oligonucleotide dissociation.
4.5

Conclusions
This work provides a detailed description and quantitative characterization of theovertone laser infrared-absorption heating method for applications in single molecule
fluorescence studies, which highlight several key advantages over conventional bulk heating
microscopy methods. First of all, the technique provides controlled and highly reproducible
experimental access to sample temperatures from 20 °C up to the boiling point (ΔT ≈ 80 °C),greatly exceeding the operating temperature ranges accessible by conventional stage-objective
heating. Secondly, use of high quality TEM00 mode laser sources permits exceptionally highspatial control and confinement of the heated volume, with typical sample regions on the order of
10-10 liters or less. Finally, this optical laser-based infrared heating system is able to achieve
controlled rises in sample temperatures with a half-maximal heating time of about 1 ms, which is
many tens of thousand times faster than the minute to hour time scales of conventional stagebased methods.
Additionally, this work demonstrates the greater utility of this optical laser heating
technique by performing pulsed heating experiments. Specifically, single-molecule dissociation
kinetics of single DNA duplexes have been studied by elevating the sample temperature by awell-characterized amount for a well-characterized duration. Most importantly, the rapid heating
and cooling properties of the system offer the ability to access kinetics at high sample
temperatures without the need for temperature-stable fluorescent dyes. These temperature
cycling dissociation studies provide useful kinetic insight into the mechanism by which a 21 bp

85

DNA duplex undergoes dissociation. In particular, the kinetic data reveal a clear sigmoidal time
lag in the appearance of the dissociated duplex. This is qualitatively inconsistent with any single
exponential rate process and indeed is more consistent with n ≈ 4 sequential first order pathways.
The current working model for this behavior is that DNA dissociation over this range of duplex
lengths is constrained by the helical twist, which makes dissociation occur from a short interior
region resulting from rapid “fraying” at both ends toward the middle. Specifically, fraying
inward with two separate rate processes from each end is shown to be six-fold more probablethan all four processes occurring from a single end, with rapid dissociation occurring when the
duplex is sufficiently frayed from both ends. In summary, the results of this study provide a“proof-of-principle” demonstration for pulsed laser heating studies of single-molecule kinetics
with high spatial (< 10-10 liter) and temporal (≈ 1 ms) control of sample temperatures, as well as
help elucidate the role of sequential, first order kinetic pathways in the dissociation of DNA
duplexes.
4.6

Experimental Methods

4.6.1

Fluorescent Oligonucleotides
A synthetically-modified, double-stranded DNA construct is purchased from acommercial supplier. The construct consists of two oligonucleotides. The first oligonucleotide(reporter strand) sequence is: 5'-|Cy3| CCC TTG |(PEG)18| CAA GGG CTC AGA CGA GTG|internal Cy5| AAA AAA AAA AAA |Biotin|-3'. The second oligonucleotide (splint strand)
sequence is: 5'-CAC TCG TCT GAG CCC TTG CAA-3'. The 36 nt reporter strand is annealed to
the 21 nt splint strand that is complementary with nucleotides 4-24 of the reporter strand,
forming a continuous 21 bp DNA duplex (Figure 4.5a).

86

4.6.2

Sample Preparation
All experiments are performed in 1-2 mm wide channels created by two pieces of 85 µm

thick double-sided sticky tape between a 170 µm thick glass coverslip and a 1 mm thick glass
microscope slide. Samples containing 575 nM Rhodamine B in water are used to measure thetemperature in all experiments designed to characterize the infrared-heating system.
4.6.3

Fluorescence Microscope
All fluorescence data are acquired on a previously described inverted confocal

microscope(58). Briefly, a pulsed (10 ps, 20 MHz) excitation source (532 nm) is directed into theback aperture of a 1.2 NA water-immersion objective via a dichroic mirror, with the resulting
fluorescence emission collected from the diffraction-limited excitation volume by the same
objective. The fluorescence photons are then spatially separated by dichroics and polarization
beam splitters onto four single-photon avalanche photodiodes (APDs), where a time-correlated
single-photon counting (TCSPC) module records the arrival time of each photon with respect toboth the most recent laser pulse (microtime) as well as the start of the experiment (macrotime).
The microtime data are used for fluorescence lifetime measurements, while the macrotime data
permit one to reconstruct time trajectories of fluorescence intensity, EFRET, lifetime, andtemperature with arbitrary choice of temporal binning.
4.6.4

Duplex Dissociation Experiments
For all duplex dissociation experiments the pre-annealed reporter-splint DNA duplex isimmobilized to the glass surfaces using previously establish biotin-streptavidin surface
immobilization techniques. The PCD/PCA oxygen-scavenging system is used to increase thephotophysical stability of the Cy3/Cy5 FRET pair(59). All experiments are performed at room
temperature (≈ 23.2 °C) in a buffer containing: 50 mM hemisodium HEPES, 100 mM NaCl, 10087µM EDTA, 2.0 mM TROLOX, 5.0 mM PCA, 100 nM PCD, 10 mM KOH, pH 7.5. Duplex
dissociation data is collected using a second excitation source to perform Alternating Laser
EXcitation (ALEX) imaging(60,61). Briefly, a red (636 nm) pulsed excitation source triggers on
the green (532 nm) 20 MHz primary excitation source. A 25 ns delay places the red laser pulses
exactly half way between green laser pulses. ALEX provides the ability to filter molecules byfluorescence, in order to ensure that only data obtained from dually-labeled fluorescent DNAs
are considered in the analysis.

88

4.7

References

1.

Joo, C.; H. Balci; Y. Ishitsuka; C. Buranachai; T. Ha. Advances in single-molecule
fluorescence methods for molecular biology, Annu. Rev. Biochem., 2008, 77, 51-76.

2.

Roy, R.; S. Hohng; T. Ha. A Practical Guide to Single-Molecule FRET, Nat. Methods,
2008, 5, 507-516.

3.

Zhao, R.; D. Rueda. RNA folding dynamics by single-molecule fluorescence resonanceenergy transfer, Methods, 2009, 49, 112-117.

4.

Holmstrom, E. D.; J. L. Fiore; D. J. Nesbitt. Thermodynamic origins of monovalentfacilitated RNA folding, Biochemistry, 2012, 51, 3732-3743.

5.

Tarsa, P. B.; R. R. Brau; M. Barch; J. M. Ferrer; Y. Freyzon; P. Matsudaira; M. J. Lang.
Detecting force-induced molecular transitions with fluorescence resonant energy transfer,
Angew. Chem. Int. Ed., 2007, 46, 1999-2001.

6.

Shroff, H.; B. M. Reinhard; M. Siu; H. Agarwal; A. Spakowitz; J. Liphardt.
Biocompatible force sensor with optical readout and dimensions of 6 nm(3), Nano Lett.,
2005, 5, 1509-1514.

7.

Fiore, J. L.; B. Kraemer; F. Koberling; R. Erdmann; D. J. Nesbitt. Enthalpy-Driven RNA
Folding: Single-Molecule Thermodynamics of Tetraloop–Receptor Tertiary Interaction,
Biochemistry, 2009, 48, 2550–2558.

8.

Graneli, A.; C. C. Yeykal; R. B. Robertson; E. C. Greene. Long-distance lateral diffusionof human Rad51 on double-stranded DNA, Proc. Natl. Acad. Sci. USA, 2006, 103, 12211226.

9.

Fiore, J. L.; E. D. Holmstrom; D. J. Nesbitt. Entropic Origin of Mg2+-Facilitated RNA
Folding, Proc. Natl. Acad. Sci. USA, 2012, 109, 2902-2907.

10.

Zhao, R.; M. Marshall; E. A. Aleman; R. Lamichhane; A. Feig; D. Rueda. Laser-assisted
single-molecule refolding (LASR), Biophys. J., 2010, 99, 1925-1931.

11.

Gross, P.; G. Farge; E. J. G. Peterman; G. J. L. Wuite. Combining optical tweezers,
single-molecule fluorescence microscopy, and microfluidics for studies of DNA-protein
interactions, Method Enzymol., 2010, 475, 427-453.

12.

Holmstrom, E. D.; D. J. Nesbitt. Real-time infrared overtone laser control of temperaturein picoliter H2O samples: "Nanobathtubs" for single-molecule microscopy, J. Phys.
Chem. Lett., 2010, 1, 2264–2268.

13.

Braun, D.; A. Libchaber. Trapping of DNA by thermophoretic depletion and convection,
Phys. Rev. Lett., 2002, 89, 188103.

89

14.

Slyadnev, M. N.; Y. Tanaka; M. Tokeshi; T. Kitamori. Photothermal temperature control
of a chemical reaction on a microchip using an infrared diode laser, Anal. Chem., 2001,
73, 4037–4044.

15.

Mao, H. B.; J. R. Arias-Gonzalez; S. B. Smith; I. Tinoco; C. Bustamante. Temperature
control methods in a laser tweezers system, Biophys. J., 2005, 89, 1308–1316.

16.

Peterman, E. J. G.; F. Gittes; C. F. Schmidt. Laser-induced heating in optical traps,
Biophys. J., 2003, 84, 1308–1316.

17.

Zondervan, R.; F. Kulzer; H. van der Meer; J. A. Disselhorst; M. Orrit. Laser-driven
microsecond temperature cycles analyzed by fluorescence polarization microscopy,
Biophys. J., 2006, 90, 2958-2969.

18.

Yuan, H.; T. Xia; B. Schuler; M. Orrit. Temperature-cycle single-molecule FRET
microscopy on polyprolines, Phys. Chem. Chem. Phys., 2011, 13, 1762-1769.

19.

Wieliczka, D. M.; S. S. Weng; M. R. Querry. Wedge shaped cell for highly absorbent
liquids: Infrared optical constants of water, Appl. Opt., 1989, 28, 1714–1719.

20.

Hung, M. S.; O. Kurosawa; M. Washizu. Single DNA molecule denaturation using laserinduced heating, Mol. Cell. Probes, 2012, 26, 107-112.

21.

Holmberg, A.; A. Blomstergren; O. Nord; M. Lukacs; J. Lundeberg; M. Uhlen. The
biotin-streptavidin interaction can be reversibly broken using water at elevatedtemperatures, Electrophoresis, 2005, 26, 501-510.

22.

Sanborn, M. E.; B. K. Connolly; K. Gurunathan; M. Levitus. Fluorescence properties andphotophysics of the sulfoindocyanine Cy3 linked covalently to DNA, J. Phys. Chem. B.,
2007, 111, 11064-11074.

23.

Marras, S. A. Selection of fluorophore and quencher pairs for fluorescent nucleic acidhybridization probes, Methods Mol. Biol., 2006, 335, 3-16.

24.

Hyre, D. E.; I. Le Trong; E. A. Merritt; J. F. Eccleston; N. M. Green; R. E. Stenkamp; P.
S. Stayton. Cooperative hydrogen bond interactions in the streptavidin-biotin system,
Protein Sci., 2006, 15, 459-467.

25.

Selvin, P. R.; T. Ha, (Eds.) (2008) Single-molecule Techniques: A Laboratory Manual,
Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.

26.

Lamichhane, R.; A. Solem; W. Black; D. Rueda. Single-molecule FRET of proteinnucleic acid and protein-protein complexes: Surface passivation and immobilization,
Methods, 2010, 52, 192-200.

27.

Akabayov, B.; A. Henn; G. Nautrup-Pedersen; M. Elbaum; I. Sagi. (2002)
Immobilization of RNA: application to single molecule spectroscopy, In Molecular,

90

Cellular and Tissue Engineering, 2002. Proc. IEEE-EMBS Special Topic Conference, pp71-72.
28.

Janissen, R.; L. Oberbarnscheidt; F. Oesterhelt. Optimized straight forward procedure forcovalent surface immobilization of different biomolecules for single molecule
applications, Colloid. Surface B., 2009, 71, 200-207.

29.

Aleman, E. A.; H. S. Pedini; D. Rueda. Covalent-bond-based immobilization approaches
for single-molecule fluorescence, ChemBioChem, 2009, 10, 2862-2866.

30.

Schwartz, J. J.; S. Stavrakis; S. R. Quake. Colloidal lenses allow high-temperature singlemolecule imaging and improve fluorophore photostability, Nat. Nanotechnol., 2010, 5,
127-132.

31.

Altman, R. B.; D. S. Terry; Z. Zhou; Q. Zheng; P. Geggier; R. A. Kolster; Y. Zhao; J. A.
Javitch; J. D. Warren; S. C. Blanchard. Cyanine fluorophore derivatives with enhanced
photostability, Nat. Methods, 2012, 9, 68-71.

32.

Chen, Y. Y.; A. W. Wood. Application of a temperature-dependent fluorescent dye(Rhodamine B) to the measurement of radiofrequency radiation-induced temperaturechanges in biological samples, Bioelectromagnetics, 2009, 30, 583-590.

33.

Karstens, T.; K. Kobs. Rhodamine B and rhodamine 101 as reference substances forfluorescence quantum yield measurements, J. Phys. Chem., 1980, 84, 1871–1872.

34.

Casey, K. G.; E. L. Quitevis. Effect of solvent polarity on nonradiative processes inxanthene dyes: Rhodamine B in normal alcohols, J. Phys. Chem., 1988, 92, 6590.

35.

Maus, M.; M. Cotlet; J. Hofkens; T. Gensch; F. C. De Schryver; J. Schaffer; C. A. M.
Seidel. An experimental comparison of the maximum likelihood estimation and nonlinear
least squares fluorescence lifetime analysis of single molecules, Anal. Chem., 2001, 73,
2078-2086.

36.

Nesbitt, D. J.; N. F. Dupuis; E. D. Holstrom. Single Molecule Studies of StreptavidinBiotin Dissociation at Elevated Temperatures, Biophys. J., 2012, 102, 270a.

37.

Lucius, A. L.; N. K. Maluf; C. J. Fischer; T. M. Lohman. General methods for analysis ofsequential "n-step" kinetic mechanisms: application to single turnover kinetics ofhelicase-catalyzed DNA unwinding, Biophys. J., 2003, 85, 2224-2239.

38.

Park, J.; S. Myong; A. Niedziela-Majka; K. S. Lee; J. Yu; T. M. Lohman; T. Ha. PCRA
helicase dismantles RecA filaments by reeling in DNA in uniform steps, Cell, 2010, 142,544-555.

39.

Ragunathan, K.; C. Joo; T. Ha. Real-time observation of strand exchange reaction withhigh spatiotemporal resolution, Structure, 2011, 19, 1064-1073.

91

40.

Watson, J. D.; F. H. C. Crick. Molecular structure of nucleic acids - a structure fordeoxyribose nucleic acid, Nature, 1953, 171, 737-738.

41.

Porschke, D.; O. C. Uhlenbeck; F. H. Martin. Thermodynamics and kinetics of helix-coiltransition of oligomers containing GC base pairs, Biopolymers, 1973, 12, 1313-1335.

42.

Dewey, T. G.; D. H. Turner. Laser temperature jump study of solvent effects onpoly(adenylic acid) stacking, Biochemistry, 1980, 19, 1681-1685.

43.

Freier, S. M.; K. O. Hill; T. G. Dewey; L. A. Marky; K. J. Breslauer; D. H. Turner.
Solvent Effects on the Kinetics and Thermodynamics of Stacking in Poly(Cytidylic
Acid), Biochemistry, 1981, 20, 1419-1426.

44.

Dewey, T. G.; D. H. Turner. Laser temperature-jump study of stacking in adenylic acidpolymers, Biochemistry, 1979, 18, 5757-5762.

45.

Ikuta, S.; K. Takagi; R. B. Wallace; K. Itakura. Dissociation kinetics of 19 base paired
oligonucleotide-DNA duplexes containing different single mismatched base pairs,
Nucleic Acids Res., 1987, 15, 797-811.

46.

Sauer-Budge, A. F.; J. A. Nyamwanda; D. K. Lubensky; D. Branton. Unzipping kineticsof double-stranded DNA in a nanopore, Phys. Rev. Lett., 2003, 90, 238101.

47.

Liphardt, J.; B. Onoa; S. B. Smith; I. Tinoco, Jr.; C. Bustamante. Reversible unfolding of
single RNA molecules by mechanical force, Science, 2001, 292, 733-737.

48.

Biancaniello, P. L.; A. J. Kim; J. C. Crocker. Long-time stretched exponential kinetics insingle DNA duplex dissociation, Biophys. J., 2008, 94, 891-896.

49.

Gunnarsson, A.; P. Jonsson; V. P. Zhdanov; F. Hook. Kinetic and thermodynamic
characterization of single-mismatch discrimination using single-molecule imaging,
Nucleic Acids Res., 2009, 37, e99.

50.

Tawa, K.; W. Knoll. Mismatching base-pair dependence of the kinetics of DNA-DNA
hybridization studied by surface plasmon fluorescence spectroscopy, Nucleic Acids Res.,
2004, 32, 2372-2377.

51.

Sinden, R. R. (1994) DNA Structure and Function, Academic Press, San Diego, CA.

52.

Yakovchuk, P.; E. Protozanova; M. D. Frank-Kamenetskii. Base-stacking and basepairing contributions into thermal stability of the DNA double helix, Nucleic Acids Res.,
2006, 34, 564-574.

53.

Porschke, D. A direct measurement of the unzippering rate of a nucleic acid double helix,
Biophys. Chem., 1974, 2, 97-101.

92

54.

Nikolova, E. N.; H. M. Al-Hashimi. Preparation, resonance assignment, and preliminary
dynamics characterization of residue specific 13C/15N-labeled elongated DNA for the
study of sequence-directed dynamics by NMR, J. Biomol. NMR, 2009, 45, 9-16.

55.

Porschke, D. Model calculations on the kinetics of oligonucleotide double helix coiltransitions. Evidence for a fast chain sliding reaction, Biophys. Chem., 1974, 2, 83-96.

56.

Dupuis, N. F.; E. D. Holmstrom; D. J. Nesbitt. Single-Molecule Kinetics Reveal CationPromoted DNA Duplex Formation Through Ordering of Single-Stranded Helices,
Biophys. J., 2013, 105, 756-766.

57.

SantaLucia, J. A unified view of polymer, dumbbell, and oligonucleotide DNA nearestneighbor thermodynamics, Proc. Natl. Acad. Sci. USA, 1998, 95, 1460-1465.

58.

Hodak, J. H.; C. D. Downey; J. L. Fiore; A. Pardi; D. J. Nesbitt. Docking kinetics andequilibrium of a GAAA tetraloop-receptor motif probed by single-molecule FRET, Proc.
Natl. Acad. Sci. USA, 2005, 102, 10505–10510.

59.

Aitken, C. E.; R. A. Marshall; J. D. Puglisi. An Oxygen Scavenging System for
Improvement of Dye Stability in Single-Molecule Fluorescence Experiments, Biophys. J.,
2008, 94, 1826-1835.

60.

Kapanidis, A. N.; T. A. Laurence; N. K. Lee; E. Margeat; X. Kong; S. Weiss.
Alternating-laser excitation of single molecules, Acc. Chem. Res., 2005, 38, 523-533.

61.

Lee, N. K.; A. N. Kapanidis; Y. Wang; X. Michalet; J. Mukhopadhyay; R. H. Ebright; S.
Weiss. Accurate FRET Measurements within Single Diffusing Biomolecules using
Alternating-Laser Excitation, Biophys. J., 2005, 88, 2939-2953.

93Chapter 5

Thermodynamic Origins of Monovalent-Facilitated RNA Folding*

5.1

Introduction
Over the last ten years, the number of structures and functions attributed to specific non-coding RNA sequences has grown significantly(1) and radically advanced the field of RNA
biology. These structures are the result of a number of smaller molecular interactions (e.g.,
tertiary interactions), which stabilize the global conformation of RNAs(2). However, it isimportant to note that these structures, although extremely helpful, represent only static imagesof RNA in a near minimum energy conformation. Under more physiological conditions, thesemolecules can be richly dynamic and occupy a broad ensemble of thermally accessible
structures(3-5). The conformational dynamics of RNA tertiary interactions contribute to thediverse array of functionality(6).
Since RNA molecules are anionic biopolymers, they are always accompanied withcounterions in solution regardless of conformation. These counterions interact favorably with theelectrostatic potential of the RNA giving rise to an “ion atmosphere.” For RNA to achieve a
specific three-dimensional conformation it must undergo any number of structural transitions,most of which result in a more compact structure. Additional cation uptake into the ion
atmosphere can help alleviate the enhanced intramolecular columbic repulsion associated with amore compact conformation. A number of studies suggest that entropic considerations arelargely responsible for cation-facilitated stabilization of folded structures(7-9). One explanation*

Adapted from: Holmstrom, Erik D; Fiore, Julie L; Nesbitt, David J. Biochemistry. 2012, 51,
3732-3743
94for this entropic promotion is as follows. As an RNA undergoes a structural transition to a morecharge-dense state, additional counterions are brought from the bulk solution to the pre-existing
ion atmosphere as a result of the elevated charge density. This restricts the translational freedomof those ions and thus gives rise to an entropic penalty associated with “unmixing” the solution.
At higher bulk concentrations, it is less entropically costly to unmix counterions that were taken
up during the folding transition, thus reducing the entropic cost of the folding transition. Singlemolecule experiments provide a relatively unexplored technique that can be used toquantitatively measure the salt-dependence of RNA folding thermodynamics and represents the
major focus of this work.
The cytosol contains cations of multiple valencies, each of which can interact with theelectrostatic potential of the RNA and thus facilitate folding(4,10-12). A number of RNAs arecapable of adopting stable, native, tertiary structures in solutions consisting solely of monovalentions(13-16). Focusing on monovalent-facilitated folding provides a crucial conceptual
simplification for both experiment and theory by avoiding complications associated with mixed
solutions of monovalent, divalent, and higher valence cations. In turn, these simple models serveas critical benchmarks for accurately testing our ability to predict thermodynamics of RNA
folding under realistic cellular conditions.
Much work has been directed at understanding the role that monovalent ions play in thefolding of structured RNAs(17-20). Generally, monovalents are capable of interacting with
RNAs through a variety of distance-dependent modes, ranging from totally diffuse to completely
chelated. The most direct mode of interaction is chelation of an ion(21,22), whereby the ion loses
multiple water molecules in its primary sphere of hydration in order to make direct, stabilizing
contact with the electronegative atoms of the RNA. These specific ion interactions are not95

dominant and therefore have a minimal effect on net charge neutralization for a single RNA in
solution(23), though with potentially more significant effects on enzymatic activity(21). Diffuse
ions have the most indirect mode of interaction with RNA. These ions interact only with theelectrostatic potential of the RNA giving rise to the above-mentioned ion atmosphere(24). Of
course there exists many other ways in which ions can interact with the RNA (e.g., indirecthydrogen bonding via the hydration layers associated with the solvated ion), but all of them fall
between the extremes of chelated and diffuse ions. This work contributes to the understanding of
how the various monovalent cation-RNA interactions alter the thermodynamics associated withformation of an isolated tetraloop−receptor tertiary interaction.
5.2

Molecular ModelRNAs are well known to fold in a hierarchical manner, with secondary structure forming

on a relatively rapid time scale (ms to ms), followed by slower folding of the tertiary structure(ms to s). Thus, a commonly proposed framework in which to view equilibrium folding of RNA
is: (25,26)
U⇔ I⇔ F
where the unfolded state (U) corresponds to an RNA without extensive secondary interactions,and the intermediate state (I) describes an RNA with well formed secondary structure(27). The I
state then proceeds to form tertiary interactions on much longer time scales, resulting in the final
folded state, F. The present work focuses exclusively on tertiary interactions by making two
additional assumptions: (i) secondary structure formation (U ⇔ I ) is rapid relative to formationof tertiary contacts and (ii) experimental conditions are such that the equilibrium constant
between U and I is much greater than unity. These assumptions allow for further simplificationof the kinetic framework to

96

I⇔ F
where the equilibrium constant (Keq) is defined as the ratio of kfold (I ⇒ F) to kunfold (I ⇐ F). In
this two-state equilibrium, both the macroscopic states I and F are likely composed of an
ensemble of microscopic states. The Boltzmann-weighted averages of the ensembles are used todescribe the macroscopic states, thus allowing for interpretations based on the two-state
unimolecular equilibrium. This simplified two-state model proves sufficient to describe theformation of an individual tertiary interaction for the RNA construct in the present study.
The model RNA used to explore tertiary interactions in monovalent ion atmospheres is asynthetic RNA construct that isolates the GAAA tetraloop−receptor interaction (Figure 5.1)(28),as observed in the P4-P6 domain of the Tetrahymena ribozyme(29).

Figure 5.1 Schematic of the three-piece donor/acceptor label RNA construct. The isolated
GAAA tetraloop−receptor serves as a model system to investigate the thermodynamics andkinetics of monovalent-facilitated formation of tertiary interactions in structured RNAs.
The construct is built from a three-piece oligonucleotide assembly that allows the GAAA
tetraloop (green), tethered by a flexible poly-A7 linker, to freely dock/undock into the 11 nt
97

receptor domain (red). This tertiary interaction(30,31) has undergone extensive biochemical andbiophysical characterization(32-36) making it a prime RNA folding motif for detailed
kinetic/thermodynamic investigation. There also exists a significant amount of structural data forthis RNA folding motif from a variety of RNA systems(21,32,37-40). Most relevantly, thetetraloop−receptor interaction is stably formed in solutions containing only monovalent ions(10),thus eliminating the complications associated with mixed monovalent and divalent ions insolution, and therefore making it an ideal RNA model to address the effect of monovalent cations
on the formation of tertiary interactions. Additionally, the “binary” (folded/unfolded) nature ofthis tertiary interaction (Figure 5.2a) makes the isolated construct used in this study particularly
useful by eliminating potential complications due to cooperative folding effects between multiple

Figure 2
tertiary interactions.
180 mM Na+ and 21 C

10.0
8.0
6.0
4.0
2.0
0.0
1.0
0.8
0.6
0.4
0.2
0.0

folded dwell time

EFRET

kHz

a)

unfolded
dwell time

1.0

P( dwell < time)

b)

1.5

2.0

2.5

3.0

time (s)1.0
kunfold

0.8

5.7(4) s

0.6

kfold

-1

8.7(5) s

-1

0.4
0.2

Figure 2: (a) Sampl
the model construct.
the FRET trajectory
the two-state behavi(b) Cumulative distriof the dwell times deexponential behavio
extract unimolecularfolding and unfolding

0.0
0.0

0.2

0.4

0.6 0.0

0.2

0.4

0.6

time (s)Figure 5.2 (a) Sample time trajectories for the model construct. A hidden Markov fit to the
FRET trajectory is used to demonstrate the two-state behavior of folding process. (b) Cumulative
distribution function (CDF) of the dwell times demonstrates single-exponential behavior and areused to extract unimolecular rate constants for folding and unfolding.

98

Lastly, the simplicity of the construct allows for the use of synthetic modifications (e.g. biotin
and fluorescent dyes) necessary for single-molecule fluorescence resonance energy transfer(smFRET) experiments (Figure 5.1)(10,28,35,36). See Materials and Methods for details.
The smFRET experiments used throughout this work are conducted using a confocal
fluorescent microscope equipped with time-correlated, two-color detection of single photons.
This techniques provides the ability to measure, in real-time and under equilibrium conditions,the amount of time of a single molecule spends in a given conformation (Figure 5.2a)(41,42). In
the framework of the previously described two-state model, the cumulative distribution function(CDF) of durations spent in either state (τdwell) will be exponentially distributed, determined by
the characteristic decay constant kunfold or kfold (Figure 5.2b)(43-45). At a specific monovalentconcentration and temperature, therefore, smFRET can be used to measure kfold, kunfold, and Keq
for the GAAA tetraloop−receptor interaction. In the present work, analysis of the equilibrium
constants as a function of both temperature and [Na+] is used to extract standard state
thermodynamics (ΔG°, ΔH°, ΔS°) for this isolated tertiary interaction. Additionally, thetemperature dependence of the forward and reverse rate constants (kfold, kunfold) is used to obtain
information about the enthalpic (ΔH‡) and entropic (ΔS‡) components of the free energy barrier(ΔG‡) as a function of [Na+].
5.3

Results

5.3.1

Folding Kinetics and Standard State Free Energy of Folding
As shown in the titration data in Figure 5.3a, increasing [Na+] both accelerates kfold anddecelerates kunfold, which clearly favors formation of products (F) over reactants (I) and therefore
increases Keq. In terms of free energy, addition of Na+ increases favorability of forming thetertiary interaction, i.e., Δ[ΔG°] < 0 (Figure 5.3b). At 130 mM Na+, the reaction is slightly99unfavorable at (ΔG° [298 K] = 0.4(1) kcal/mol), whereas at 830 mM Na+, folding becomesappreciably favored (ΔG° [298 K] = -2.0(1) kcal/mol). Clearly, the free energy, and thereforeln[Keq], has a non-liner dependence on [Na+].

Fig. 3100

kunfold or kfold(s-1)

a)

10

1
0.1

G°  (kcal/mol)b)1[Na+] M

1
0
-1
-2
0.0

0.2

0.4

0.6

0.8

1.0

+

[Na ] mM
4ln[kfold/kunfold]

C)

3
2

170(20) mM Na+
G°  =  0  2± = 2.4(1)

1
0
-1
-2.5

-2.0

-1.5

-1.0

-0.5

ln[aNa+]Figure 5.3 (a) Na+-dependent kinetics for the formation of the GAAA tetraloop−receptor
interaction. Increasing [Na+] yields a significant enhancement of the folding rate constant (kfold)
and a subtle reduction in the rate constant for the unfolding process (kunfold). (b) The dependenceof the free energy on [Na+] and (c) preferential interaction coefficient analysis for the formationof the tertiary interaction, which yields an ion pair uptake of 2ΔΓ± = 2.4(1) and no change in freeenergy at [Na+] = 170(20) mM.

100

These data, when considered in the context of the physically-motivated theoretical frameworkprovided by preferential interaction coefficients, can be used to create a more informative
depiction of the Na+-dependence of Keq (7,46). Based on this model-free description of RNA-ion
interactions, the logarithm of the equilibrium constant (ln[Keq]) should scale linearly with thelogarithm of the thermodynamic activity of the monovalent ion (ln[aNa+]). From such ananalysis, the slope (2ΔΓ±) corresponds to the net change in the number of ion pairs associatedwith the ion atmosphere as a result of the folding transition. This is well demonstrated by a linear
fit of ln[Keq] vs ln[aNa+] where the slope yields a net uptake of approximately 2.4(1) ion pairs(Figure 5.3c). This plot also demonstrates that at 170(20) mM Na+ formation of the tertiary
interaction results in no free energy change. It should be noted that because ln[Keq] is
proportional to Gibbs free energy, ΔG° = ΔH° – TΔS°, it seems a reasonable anticipation thatboth ΔH° and TΔS° might also scale linearly with ln[aNa+].
The purpose of this work is to investigate the thermodynamics associated with theformation of the GAAA tetraloop−receptor interaction as a function of monovalent salt
concentration. Therefore, it will be useful to develop the language used to describe the range ofexperimental [Na+]. Specifically, this work will refer to RNA molecules in 180 mM Na+ as being
under “free energy neutral” conditions where ΔG° ≈ 0 kcal/mol. Progressively higher
concentrations of Na+ (i.e., 330 mM, 480 mM, and 630 mM) correspond to progressively more
“free energy favorable” conditions as ΔG° decreases below zero with addition of Na+.
5.3.2

Standard State Enthalpy and Entropy of Folding
The observed increase in kdock and decrease in kundock represent the fundamental kinetic

basis for Na+-promoted folding (Figure 5.3a). However, in order to provide a more insightfuldescription of the free energy folding pathway from I to F, it is also important to understand the101thermodynamic contributions (i.e. enthalpic and entropic) to monovalent-facilitated folding. Thiscan be best accomplished by temperature dependent van’t Hoff analysis of Keq(36), which allows
the free energy (ΔG°) to be partitioned into the enthalpy (ΔH°) and entropy (ΔS°) changes
associated with folding at a particular [Na+] (Figure 5.4a). Specifically, in a plot of ln[Keq] vs.
1/T the slope and intercept are equal to -ΔH°/R and ΔS°/R, respectively.

Fig. 4a)3ln [Keq]210-1

180 mM 330 mM
480 mM 630 mM

-2
3.25

b)

3.30
-1

3.35

3.40

3

1/T (K ) x 10

ΔH°
kcal/molΔS°
cal/mol/K

180-26(2)-86(6)

330-19(3)-60(8)

480-12(2)-36(8)

630

-4(1)

-8(4)

c)c)

0

kcal/mol

[Na+]
mM

-10
-20

ΔH°
TΔS° (298K)

-30
-2.5

-2.0

-1.5

-1.0

-0.5

ln[aNa+]Figure 5.4 (a) van’t Hoff analysis of the tetraloop−receptor interaction at four [Na+]. Increasing[Na+] increases both the ΔH° (decreasing slope) and the ΔS° (increasing vertical intercept) for
the formation of the tertiary interaction. (b) Table of thermodynamic parameters from the van’t
Hoff analysis. (c) Bar graph to visually display the change in ΔH° and ΔS° as a function ofln[aNa+].

102

Under free energy neutral conditions of 180 mM, the van’t Hoff plot in Figure 5.4a
reveals the folding process to be quite exothermic (ΔH° = -26(2) kcal/mol). This is entirely
consistent with the formation of 11 hydrogen bonds in the F state, which are absent in the I
state(37). In principle, an additional source of exothermicity could be the site-specific chelationof a monovalent ion in the folded conformation. Indeed, there is some structural evidence that
supports a proposed monovalent binding site within the tetraloop−bound receptor domain(21).
However, these chelation interactions are considerably stronger and thus likely already saturated
well below the present working conditions. Therefore, changes in [Na+] over the range of 180 to
630 mM will not significantly affect how monovalent chelation contributes to the folding
thermodynamics. Under the same free energy neutral experimental conditions, the van’t Hoffanalysis reveals a folding process that is entropically penalized (ΔS° = -86(6) cal/mol/K). This
loss of disorder associated with formation of the F state is quite reasonable given that the noncanonical hydrogen bonds within the receptor domain are more conformationally restricted after
the formation of tertiary contacts. Furthermore, formation of the tertiary interaction restricts theconformational freedom of the poly-A7 linker. Lastly, the large entropic cost to folding is also
the result of ion uptake as the RNA undergoes a structural transition. Based on theseexperimentally measured values of ΔH° and ΔS°, the calculated Gibbs free energy for the folding
process at 180 mM Na+ is nearly zero (ΔG° [298 K] = -0.4(9) kcal/mol), which is consistent withthe predictions from the preferential interaction coefficient analysis (Figure 5.3b).
These three thermodynamic parameters (i.e., ΔG°, ΔH°, ΔS°) at 180 mM now establish abaseline with respect to increasing monovalent concentrations. If the effect of Na+-facilitatedfolding were purely related to ion uptake from solution (e.g. unmixing), one would predict that103

increasing [Na+] would have no affect on the enthalpy of folding (i.e. Δ[ΔH°] = 0). Surprisingly,
both the folding entropy and enthalpy increase with increasing [Na+] (Figure 5.4). The van’t
Hoff plot for the equilibrium docking reaction at 330 mM Na+ (and higher) demonstrates a lesspositive slope (Δ[ΔH°] > 0) when compared to free energy neutral conditions. Specifically, theexothermicity of folding continues to diminish with further increases in [Na+]. Indeed, under themost free energy favorable conditions, formation of the tertiary interaction is much lessexothermic (ΔH° =-4(1) kcal/mol). A similar and slightly over-compensating trend is also
apparent when probing changes in the standard state entropy of folding (i.e., intercepts of the
van’t Hoff plot) as a function of [Na+]. Increasing [Na+] gives rise to a large increase in entropychange (Δ[ΔS°] > 0) for folding. At 630 mM Na+, the entropic penalty for folding is greatly
reduced (ΔS° = -8(4) cal/mol/K). As apparent in Figure 5.4c, both ΔH° and TΔS° scale linearlywith ln[Na+].
Finally, the Van’t Hoff analysis in Figure 5.4a demonstrates that increasing [Na+]systematically increases both ΔH° and ΔS°, but that at a fixed temperature, increasing [Na+]increases the equilibrium constant and thus decreases ΔG°. In order to have decreasing freeenergy (Δ[ΔG°] < 0) with increasing [Na+], the rate of increase for TΔS° must be larger than therate of increase for ΔH° (Figure 5.4c). This implies that an increase in monovalent cation
concentration enhances tetraloop−receptor folding via a reduction of the entropic penalty thatslightly over-compensates for the loss in exothermicity. Stated simply, the well-known increasein stability of the tetraloop−receptor tertiary interaction with increasing monovalent cation
concentration is in fact the result of entropic considerations. It is also interesting to note thatrather large changes in ΔH° and ΔS° give rise to relatively small changes in ΔG°. This nearly
compensatory nature of Δ[ΔH°] and Δ[ΔS°] is intriguing and may provide a way to tune the104temperature sensitivity of biochemically relevant processes (e.g., RNA folding) without
significantly perturbing the equilibrium constant.
5.3.3

Transition State Thermodynamics
This trend of increasing ΔH° and ΔS° with increasing [Na+] clearly provides surprisinginformation about thermodynamic changes between reactants (I) and products (F). Taken one
step further, a parallel investigation of the transition state barrier can provide additional insight
into how the monovalent ion atmosphere affects the GAAA tetraloop−receptor interaction.
Specifically, the temperature dependence of kfold and kunfold can be analyzed in the context oftransition state theory (TST) to provide thermodynamic details about access to the transition statefrom the intermediate (I) and folded (F) states, respectively(47,48). Although more sophisticatedtreatments exist in the literature(49), for present purposes we consider TST in its simplest formof activated complex theory. This treatment invokes two key assumptions: (i) a well definedtransition state dividing surface, beyond which the reactants proceed toward products without
recrossing and (ii) a transition state in quasi-equilibrium with the reactants. The transition state
expression for the folding rate constant can then be rigorously written as‡(q perp )‡- ΔU foldkT
kT
k fold = be bIIh (q perp )(q para )(Eq. 5.1)

where kb, T, and h are Boltzmann’s constant, temperature and Planck’s constant respectively.
In Eq. 5.1,

qIperpandqIpararepresent partition functions for the I state in all degrees of‡

freedom (i.e., both perpendicular and parallel to the reaction coordinate), whereas

q perprepresents the corresponding partition function at the transition state, including only degrees of
freedom perpendicular to the reaction coordinate. Additionally, ΔU‡fold corresponds to the105potential energy barrier between the I state and the transition state. Eq. 5.1 can be conveniently
expressed in terms of the Gibbs free energy barrier (ΔG‡fold) by rewriting the ratio of the partition
functions associated with the perpendicular degrees of freedom as Eq 5.2.
‡

(q perp )
(q

I
perp

)

ΔS ‡foldkT=e b

If pressure-volume work is assumed to be negligible (i.e., ΔH = ΔU) and(Eq. 5.2)

qIparato be in the hightemperature limit (i.e., kbT/hn), the TST equation reduces to the simple activated complex
expression:‡- ΔG fold

k fold =ν e kbT(Eq. 5.3)

where n corresponds to the attempt frequency—parallel to the reaction coordinate—for
approaching the transition state. Eq. 5.3 can be readily expressed as an Eyring plot, ln[kfold/ν] vs
1/T , whose slope and intercept correspond to the transition state enthalpy (-ΔH‡fold/R) andentropy (ΔS‡fold/R), respectively. Though commonly misinterpreted as such, note that the attempt
frequency (υ) is not the same as kbT/h, and in fact an accurate determination of n requires far
more detailed knowledge of the true folding potential surface than currently available. The
present analysis uses υ ≈1×1013 sec-1 as an approximate order of magnitude for hydrogen bondformation/breaking rates. However, it is important to stress that the choice of ν only introduces aconstant offset to the barrier entropy (ΔS‡fold) and has no effect on the barrier enthalpy (ΔH‡fold).
Furthermore, the choice of n also has no effect on the change in barrier entropy (Δ[ΔS‡fold]) as afunction of [Na+], which is of particular interest for the analysis and interpretation of the present
results.

106

5.3.4Temperature Dependence of kfold
An Eyring plot of the folding rate constants (Figure 5.5a) at 180 mM Na+ depicts aslightly positive slope, indicating that formation of the transition state under free energy neutral
conditions is slightly exothermic (ΔH‡fold = -3(2) kcal/mol), which is consistent with recent
studies of a similar RNA construct under similar conditions(50).

Fig. 5ln [kfold/]a) -25-26-27-28

180 mM 330 mM 480 mM 630 mM
3.25

3.30
-1

3.35

3.40

3

1/T (K ) x 10

b)

[Na+]
mM

ΔH‡fold
kcal/mol

ΔS‡fold
cal/mol/K

180

-3(2)

-64(7)

330

0.3(7)-53(2)480

5(1)-35(3)

630

7.7(8)-25(3)

kcal/mol

c) 10
0
-10

ΔH‡

-20

TΔS‡ (298K)
-2.5

-2.0

-1.5

-1.0

-0.5

ln[aNa+]Figure 5.5 (a) Transition state theory (TST) analysis of the rate constant associated withformation of the tetraloop−receptor interaction (kfold) as a function of temperature. (b) Table ofthermodynamics parameters from the TST analysis. (c) Bar graph depicting the change in ΔH‡fold
and ΔS‡fold at four values of ln[aNa+].

107

The lack of an endothermic component of the free energy barrier suggests the transition state is
structurally similar to the I state, i.e., the transition state is “early” with respect to reactants.
Again, if monovalent-facilitated folding were solely a result of entropic terms associated with
unmixing of the solution, one would expect the enthalpic component of the free energy barrier tobe insensitive to [Na+]. Surprisingly, for increasing [Na+], the slope of the Eyring plot becomessignificantly more negative (Δ[ΔH‡fold] > 0), indicating the emergence of an endothermic barrier,which is often associated with formation of a less early transition state. This is best demonstrated
at 630 mM Na+, where the enthalpy for forming the transition state is significantly positive(ΔH‡fold = 7.7(8) kcal/mol). One source of an endothermic folding barrier at elevated [Na+] could
be associated with preferential stabilization of non-native contacts that must break prior toforming the transition state.
Additionally, the intercepts of the Eyring plot (Figure 5.5a) demonstrate that the entropychange associated with formation of the transition state also increases (Δ[ΔS‡fold] > 0) with [Na+].
For example, at 180 mM Na+, transition state formation is a heavily penalized process (ΔS‡fold = 64(7) cal/mol/K). By 630 mM Na+, however, this penalty is dramatically reduced (ΔS‡fold = 25(3) cal/mol/K). These combined results suggest that: (i) folding under free energy neutral
conditions is predominantly rate-limited by a conformational search (e.g., -TΔS‡fold > ΔH‡fold ≈ 0
kcal/mol) and (ii) while under most free energy favorable conditions, folding is rate-limited byan increased amount of structural rearrangements and reduced conformational search (e.g., TΔS‡fold ≈ ΔH‡fold > 0kcal/mol).Again, it should be noted that both ΔH‡fold and TΔS‡fold appear toscale linearly with the ln[Na+] (Figure 5.5c).
Based on the simple relationship between Gibbs free energy, entropy, and enthalpy, we
can now describe the free energy barrier as a function of monovalent ion concentration.

108

Increasing the folding favorability of the experimental conditions gives rise to a strong increasein the entropic and enthalpic components of the free energy barrier. Similar to what was noted
above for the standard state folding thermodynamics, these large changes yield a much more
subtle effect on the transition state free energy barrier. Specifically, ΔG‡fold [298 K] decreases
only by ≈ 0.9 kcal/mol (from 16.1(8) kcal/mol to 15.2(7) kcal/mol) between free energy neutral
and the most free energy favorable conditions. This drop in barrier height is the result of adramatic increase in entropic rewards (≈ 11.6 kcal/mol at 298 K) that slightly outweighs theincrease in enthalpic cost (≈ 10.7 kcal/mol) associated with formation of the transition state (i.e.,
Δ[TΔS‡fold] > Δ[ΔH‡fold]). As a central result, this implies that the Na+-accelerated kfold of the
GAAA tetraloop−receptor interaction (Figure 5.3a) occurs via a subtle reduction of the folding
free energy barrier, which results from entropic facilitation barely overcoming enthalpic
suppression.
In order to determine if the poly-A7 linker has any appreciable effect on thethermodynamics of tetraloop−receptor folding, a poly-U7 linker is used to replace the poly-A7
linker of the original construct (Supplemental Figure S5.1). As was the case with the originalpoly-A7 construct, addition of Na+ resulted in an acceleration of kfold and a deceleration of kunfold.
A simple van’t Hoff analysis (Supplemental Figure S5.2) with the poly-U7 RNA constructreveals the same trends as the poly-A7 RNA construct, i.e., increasing [Na+] from 80 to 330 mM
increases ΔH° from -26 to -13 kcal/mol. This reduction of exothermicity was subtly
overcompensated by an increase in ΔS° from -88 to -38cal/mol/K. Furthermore, a corresponding
Eyring analysis of the forward rate constant indicates that increasing [Na+] also increases both

ΔH‡fold and ΔS‡fold (Supplemental Figure S5.3)—the same trend observed for the poly-A7
construct. The additional analysis of the poly-U7 construct suggests that, although the linker may

109subtly change the thermodynamics of folding at a given [Na+], the observed trends in the A7-

Supplemental Figure 1

linker based construct are dominated by the GAAA tetraloop−receptor tertiary interaction.

U7 ConstructSupplemental Figure 1
three-piece donor/accep
construct. The poly-U7 li
determine the effect of th
folding free energy barrie

Supplemental Figure S5.1 Schematic of the three-piece donor/acceptor label RNA construct.
The poly-U7 linker is used to determine the effect of the linker on the folding thermodynamics.

110

upplemental Figure 2a)

U7 Construct
[Na+]
mM

ΔH°fold
kcal/mol

ΔS°fold
kcal/mol/K

080-26(1)-88(4)-1100-23.9(9)-80(3)

80 mM 100 mM
180 mM 330 mM

180-22.0(9)-71(3)

3.35

330-13(2)-38(6)b)3

ln [Keq]

2
1

-2
-3
3.25

3.30
-1

3.40

3

-20
-30
100

200

(kcal/mol/K)

d)

-10

S°

H°

c)

(kcal/mol)

1/T (K ) x 10

-30
-60
-90
100

300

200

300

[Na+]

[Na+]

Supplemental Figure S5.2 (a) van’t Hoff analysis of tetraloop−receptor interaction at four [Na+]with the poly-U
Increasing[Na+] increases
both theΔH° (decreasing
and the ΔS
7 linker.2:
Supplemental
Figure(a) Transition
state
theory(TST)slope)
analysisof° the rate
(increasing vertical intercept) for the formation of the tertiary interaction. (b) Table ofconstantthermodynamic
associatedwith formation
tetraloop—receptor
interaction
from the van’t ofHofftheanalysis.
(c) Visual display of changes
in ΔH°
upplemental
Figure 3parameters+
U
Construct
and
ΔS
°
as
a
function
of
ln[Na
].
(k ) as a function of temperature7 with the poly-U linker based construct (b)
7

fold

-25
-26

mM

kcal/molkcal/mol/K

80

-4(1)

-69(3)

100-3.3(4)-65(1)

180-0.7(3)-54(1)

330

8.7(1)-21(5)

-27
-28
-29

80 mM 100 mM 180 mM 330 mM
3.25

3.30

3.35
-1

3.40

3

0
-10
100

200

[Na+]

300

(kcal/mol/K)

d)10

S‡fold(kcal/mol)

1/T (K ) x 10c)H‡fold

ln [kfold/]Table of thermodynamics parameters from the TST analysis (c,d) Bar graph
‡
‡
[Na+]
-24
depicting a)thechange in ΔH° and ΔS° atb)four
valuesΔH
of fold[Na+] ΔS fold

0
-20
-40
-60
-80
100

200

300

[Na+]

Supplemental Figure S5.3 (a) Transition state theory (TST) analysis of the rate constantassociated with formation of the tetraloop−receptor interaction (kfold) as a function of temperature
with the poly-U7 linker based construct (b) Table of thermodynamics parameters from the TST
Supplemental
Figure 3: (a) van’t Hoff analysis of tetraloop—receptor interaction
analysis (c) Bar graph depicting the change in ΔH‡fold and ΔS‡fold at four values of ln[Na+].

at four [Na+] with the poly-U7 linker. Increasing [Na+] increases both the ΔH‡(decreasing slope) and the ΔS‡ (increasing vertical intercept) for the formation of111
the tertiary interaction (b) Table of thermodynamic
parameters from the van’t Hoffanalysis (c,d) Visual display of changes in ΔH‡ and  ΔS‡ as a function of [Na+].

5.4

Discussion

5.4.1

Folding Pathways and Ion Atmospheres
The free energy barrier thermodynamic data can be used in conjunction with thepreviously extracted standard state thermodynamic data to provide additional insight into the“reaction coordinate” for tetraloop−receptor folding as a function of [Na+]. As summarized
schematically in Figure 5.6, the reaction coordinate follows the folding pathway from left toright, where the horizontal distance between the I state and the transition state qualitatively

b)

30
20
10
0
10

-T(298 K)S
(kcal/mol)a)H (kcal/mol)

represents how “early” or “late”
the free6energy barrier is along the reaction coordinate.
Figure0-10

180 mM
630 mM

-20
-30

c)
G
(kcal/mol)

20
15
10
5
0

I

TS

F

Figure 6: Comple
reaction coordinat
the GAAA tetraloointeraction. The thenthalpy, (b) entro
along the reaction
saturated” and sat
densities. The horistates qualitativelyor ‘late’ they are acoordinate. See teFigure 5.6 Complete thermodynamics reaction coordinates for the formation of the GAAA
tetraloop−receptor tertiary interaction. The three panels depict the: (a) enthalpy, (b) entropy, and(c) free energy along the reaction coordinate at 180 and 630 mM Na+. The horizontal distance
between states qualitatively represents how “early” or “late” they are along the reaction
coordinate (see Section 5.4 for details).

112

For all F states in the plot, the absolute energies are referenced to 0 kcal/mol. This choice ismade because the F state is least likely to be influenced by [Na+], and it will therefore moreclearly demonstrates changes to the folding pathway(51). First, the implications of the 180 mM
reaction coordinate will be considered.
5.4.2

Folding in a Free Energy Neutral Monovalent Ion Atmosphere
The path of the reaction coordinate under free energy neutral conditions (Fig 5.6, purple)

is easily interpreted, and serves as a baseline for comparison to the 630 mM Na+ reaction
coordinate. Formation of a number of tertiary contacts in the F state, and perhaps specificchelation of a monovalent ion, makes the folding process strongly exothermic. Furthermore, themeasured ΔH° values reported herein are in decent agreement with previously reported values
using a complementary technique(50). Specifically, the measured exothermicity of the poly-U7
construct at 180 mM Na+ is almost exactly half of published values for a dual tetraloop−receptor
construct at 200 mM KCl; the poly-A7 construct is slightly more than half as exothermic.
The absence of any significant endothermic component of the free energy barrier (i.e., anearly transition state) suggests most of the standard state enthalpic gains occur as the transition
state forms the F state (Figure 5.6a, purple), which is again supported by previous work(50). The
strong entropic component of the free energy barrier at 180 mM Na+ suggests that there is asignificant conformational search of the receptor by the tetraloop, which represents the ratelimiting step in transition state formation. Previous structural studies of the receptor domain haveidentified a high degree of conformational disorder within the free receptor, which is absent once
the full tertiary interaction has formed(37). Therefore, the loss of entropy associated with theformation of the transition state, and subsequently the F state, can in part be attributed to therestricted orientation of the base pairs within the 11-nucleotide receptor (Figure 5.6b, purple).

113

Other contribution to the loss of entropy for folding under free energy neutral conditions comefrom: (i) localizing the tetraloop near the receptor and (ii) the subsequent uptake of ions from thebulk to the ion atmosphere in order to compensate for the increased charge density of both thetransition state and the F state. This leads to a proposed compact transition state where thetetraloop is proximal to the receptor and a few additional monovalent ions have been localizedaround the RNA. Furthermore, the nucleotides within the receptor must have transiently adopted
a conformation from the ensemble of unfolded microstates that is “aligned” for the formation ofhydrogen bonds with the GAAA tetraloop.
Our identification of (i) an early transition state and (ii) a large entropic barrier in the
GAAA tetraloop−receptor interaction supports two emerging paradigm for RNA folding. First,
our findings under free energy neutral conditions support the observation that transition states for
tertiary interactions are often early (40, 47, 48,(48,52), (50)); that is to say that there is little to noenthalpic component to the free energy barrier (ΔH‡fold ≈ 0 ). Furthermore, the idea of an earlytransition state implies that the structure of the transition state more closely resembles the I statethan the F state. The second paradigm supported by the thermodynamics observations under freeenergy neutral conditions is that RNA folding is accompanied by a large entropic component of
the free energy barrier (ΔS‡fold < 0 ) resulting from a conformational search. This idea was first
proposed in regard to the slow folding of ribozymes from group I(52) and group II (53) introns,
where the notion of contact order was used to explain the large entropic barrier to folding. The
idea of a rate-limiting conformational search and a large ΔS‡fold has since been supported byexperiments on the hairpin ribozyme(44,54,55) and the GAAA tetraloop−receptor(48,50). More
importantly, however, this work provides a quantitative framework for understanding how the114

entropic and enthalpic components of the free energy barrier change with increasing monovalentcation concentrations.
5.4.3

Folding in a “Free Energy Favorable” Monovalent Ion Atmosphere
Increasing the [Na+] dramatically alters the folding pathway associated with formation of

the tertiary interaction (Figure 5.6, orange). At elevated [Na+] the standard state entropy andenthalpy for folding are significantly increased as well as the entropic and enthalpic componentsof the free energy barrier. Mg2+-induced increases to ΔS° and ΔH° been observed in both the P4P6 domain of the Tetrahymena ribozyme as well various GAAA tetraloop−receptor construct(48,56,57). However, support for the same trend with monovalent ions is less apparent in theliterature. A great deal of work has concluded that for duplex denaturation there is little to nomonovalent-dependence of ΔH°(8,58) concluding that entropy is the only thermodynamic termthat is dependent on monovalent ion concentration. However, other have seen significant
increases in ΔH° for helix formation with increasing monovalent salt concentration(59),suggesting that both entropy and enthalpy must be changing with addition of monovalent salt.
With regard to the formation of the GAAA tetraloop−receptor interaction, recent studies of abimolecular, dual tetraloop−receptor RNA construct were not able to unambiguously determine
whether the observed monovalent-induced stability was the result of entropic or enthalpic
considerations(50). However, they note that over the range of 100-300 mM KCl the Δ[ΔS°]
accounts for 134-166% of the overall Δ[ΔG°], which clearly implies that the increase in ΔH°
must be 34-66% of the overall Δ[ΔG°], which is certainly qualitatively consistent with ourobservations that both enthalpy and entropy change and that the increase in ΔS° slightly
overcomes the increase in ΔH°.

115

These results, perhaps, suggest that addition of salt may indeed increase the observed

ΔH° of formation. Although these bimolecular, dual tetraloop−receptor constructs do contain the
GAAA tetraloop−receptor interaction from the Tetrahymena ribozyme, the context of theinteraction is quite different. The duality of the construct and the significantly enhanced degreeof helical packing upon formation of the interaction make truly rigorous comparisons with theconstruct used for this study challenging at best. The degree to which ΔH° for various RNA
folding motifs is affected by monovalent salt concentrations is likely a result of how many
alternative conformations are easily accessible within the ensemble of microstates associatedwith the I state. Nevertheless, in the examples above, addition of monovalent salt increases theoverall favorability of folding by increasing the ΔS° of formation.
In conjunction with the present work, these observations support a potential paradigm
fundamental to tertiary interactions and perhaps even RNA folding, whereby additionalmonovalent cations facilitate folding by reducing the entropic penalty (Δ[ΔS°] > 0) at thepotential cost of reducing the folding exothermicity (Δ[ΔH°] > 0). The qualitatively establishedfree energy barrier thermodynamics for the GAAA tetraloop−receptor are used to further support
this paradigm by providing a physical picture based on the molecular structure of the tertiary
interaction, as described below.
It is important to remember that discussion of structural changes pertains primarily toeither of the Boltzmann-weighted macroscopic states; it is likely that these structural changes
reflect a shift in the energetic distribution of the ensemble of microscopic states. As illustrated by
the orange traces in Figures 5.6a and 5.6b, formation of the transition state in the most freeenergy favorable conditions is accompanied by an enthalpic cost and an entropic penalty. The
enthalpic cost associated with transition state formation is attributed to preferentially stabilized116

non-native intramolecular contacts within the unbound receptor domain (I) that must be broken
prior to forming the transition state. One such contact from structural models of the freeFigure 7 domain, is that of base U19 (numbering from ref (37)). An NMR solution
tetraloop−receptor
structure of the free receptor domain(37) depicts a conformation where U19 participates in one
hydrogen-bonding interaction with U5 and is basestacked between G20 and A6 (Figure 5.7).

free receptor

3

G8

U17

bound receptor

5

A7

3

kfold

A18

5

U19

C4

G20

C3

G21

kunfold
3

5

U17

5

A6

A7

A6
U5

G8

U5

A18

C4

G20

C3

G21
3

Figure 5.7 Structural representation of the free and bound receptor domain. Bases that undergo
significant structural rearrangement upon forming the tetraloop−receptor interaction are colorcoded. Bases of the GAAA tetraloop are not shown to emphasize changes within the receptor
domain (see Section 5.4.3 for details).

117

Figurerepresfree anreceptBasessignific
rearraformintetralointeraccoded
details

Structural models of the bound receptor domain show that in the presence of the GAAA
tetraloop the U19-U5 hydrogen bond and all of the basestacking interactions with U19 are absent
as U19 is flipped outside of the receptor to accommodate the tetraloop(29). The reduction of theentropic barriers results from the I state being preorganized at high [Na+] by non-native contacts,
thus removing the entropic cost associated with forming an “aligned” receptor conformation. The
remaining entropic penalty for forming the transition state at 630 mM Na+ is likely associatedwith bringing the tetraloop in close proximity to the receptor and ion uptake associated with theincreased charge density. Taken together, these conclusions can be used to portray a moredetailed image of the transition state. Specifically, the tetraloop is (i) proximal to the receptor,
which is (ii) in a conformation “aligned” for formation of the tertiary interaction and (iii) has
U19 flipped outside of the receptor to accommodate the GAAA tetraloop.
This putative model of the transition state is also in accord with the observed changes in
ΔH‡fold and ΔS‡fold with increasing [Na+]. At 180 mM Na+, the interactions involving U19 in thefree receptor are only poorly stabilized. Addition of Na+ preferentially stabilizes theseinteractions within the free receptor (I), which gives rise to an increase in ΔH‡fold associated withhydrogen-bond breakage, unbasestacking, and base flipping—all prior to forming the transition
state. The increase in ΔS‡fold comes from the fact that stabilizing the non-native interactions preorganizes the receptor, thus reducing the amount of organization (i.e., entropy loss) required toform the transition state. An additional source of increasing ΔS‡fold could be global compaction of
the RNA at elevated [Na+], which is supported by a Na+-dependent undocked FRET
position(35). This would reduce the accessible volume for the unbound tetraloop, thus making it
less entropically costly to localize the tetraloop proximal to the receptor in order to form thetransition state. As additional support for this molecular model, recent electron paramagnetic

118

resonance (EPR) experiments with spin-labeled nucleotides have demonstrated that increasing
the cation concentration by itself does not promote the unbasestacked conformation of U19(60).
Indeed, only in the presence of GAAA tetraloop is the U19 base able to adopt a stableconformation where it is flipped outside of the receptor domain, which further supports the
proposed origin of monovalent-facilitated folding of the GAAA tetraloop−receptor interaction.
The tertiary contacts within the unbound receptor associated with U19 are used to explain
the observed changes in the free energy barrier to folding. However, it is possible thatinteractions with solution may also contribute. Recent experiments have highlighted theimportance of phosphate backbone dehydration in RNA folding transitions(61). With this inmind, solvent rearrangement may serve as potential explanation for the monovalent-inducedchanges to the folding thermodynamics. However, one must have a detailed understanding forhow changing the monovalent concentration affects the entropic and enthalpic components ofphosphate backbone dehydration. Fortunately, the total amount of buried phosphate surface area
for the tetraloop−receptor interaction used in this study (≈67 Å2) is much smaller than morebiologically relevant RNAs making the energetics associated with desolvation much smaller inmagnitude than it may have otherwise been(61).
One last explanation for the strong salt-dependence of both the enthalpic and entropic
terms pertains to the polynucleotide linker used to create the unimolecular tetraloop−receptor
construct. Theoretically, increasing the monovalent concentration could preferentially stabilize arigid, base-stacked conformation within the poly-A7 linker(62). If so, elevated monovalentconcentrations could result in an additional enthalpic loss (and entropic reward), arising from theneed to disrupt this rigid base-stacked linker.

119Control experiments can be performed to test and rule out this alternative explanation.
The poly-U7 single strand RNA linker is thought to only be rigid and structured at low ionicstrengths (e.g., < 50 mM monovalent)(63) or at low temperatures (T< 10 °C)(64). As a result thepoly-U7 linker should act as a more flexible, passive, linker that is less susceptible to Na+facilitated base-stacking. A comparison of the results of the poly-U7 construct with the rigourusly
determined values of the poly-A7 construct clearly suggests the polynucleotide linker has only aminimal contribution to the absolute thermodynamic values, and that the observed changes in allmeasured thermodynamic values (ΔH°, ΔS°, ΔH‡fold, and ΔS‡fold) follow the same trend withincreasing [Na+]. One interesting observation is that at both 180 and 330 mM Na+, formation of
the GAAA tetraloop−receptor interaction is more exothermic for the poly-A7 RNA constructthan for the poly-U7 RNA construct (i.e., ΔH°[A7] < ΔH°[U7]) (Figure 5.4b and Supplemental
Figure S5.2b). This suggests that perhaps the poly-A7 linker forms and maintains a few basestacking interactions along the folding pathway. We note that the effect of single stranded linkers
and base-stacking in RNA folding is a particularly important problem for: (i) H-type pseudoknot
formation (ii) other single-stranded regions of larger functional RNAs (e.g., the templateboundary element of human telomerase RNA), and (iii) oligonucleotide duplex formation.
Consequently, future work directed at elucidating differences in folding kinetics andthermodynamics due to changes in single-strand structure would be particularly relevant.
5.4.4

Dependence on Monovalent Cation Identity
One issue with RNA folding studies is the potential role of monovalent identity. For

example, the use of K+ vs Na+ cations has been shown to affects rates of RNA catalysis, and has
been attributed to specific binding of K+(21). We can easily test for monovalent identity effects
with parallel folding studies as a function of [K+]. Interestingly, the corresponding K+ vs. Na+

120

titrations reveal completely negligible differences in the folding kinetics of the GAAA
tetraloop−receptor construct as summarized in Figure 5.8(65), which is consistent with existingstudies of a GAAA tetraloop−receptor
Figure 8 construct(18).

++

kunfold (s-1)

kfold (s-1)

Figure 8: Monovalen
dependent kinetics foNa
the formation of the
K
GAAA
14
tetraloopreceptor
1210interaction. Increasin
86
monovalent42
concentration provide
0
0
300
600
900
a significant[M ] mMenhancement of the
Figure 5.8 Monovalent-identity and folding kinetics for the formation of the GAAA
tetraloop−receptor interaction. Increasing monovalent concentration provides afolding
significant rate constantenhancement of the folding rate constant (kfold) and a subtle reduction in the rate constant for the(k K+. ) and a subtle
unfolding process (kunfold). No significant difference is observed between Na+ and foldreduction in the rate
constant
Furthermore, temperature-dependent kinetic experiments identical to those describedfor Na+ for theunfolding process
reveal that all thermodynamic parameters for Na+ and K+ promoted folding are identical within(kunfold).
70
60
50
40
30
20
10
0

+

experimental uncertainty (Figure 5.9). The insensitivity to monovalent identity suggests that theeffects observed are common to both Na+ and K+, thus demonstrating the broader applicability ofthe proposed model for monovalent-facilitated RNA folding.
5.5

Conclusion
The present work describes the use of smFRET experiments to better understand theorigins of monovalent-facilitated formation of an RNA tertiary interaction. The kinetic origin of
this enhanced folding results from a large acceleration of kfold and a subtle deceleration of kunfold
at elevated [Na+]. The large 14-fold change in the forward folding rate constant is the result of a121

with the folding reaction. This reduction in the free energy barrier is relatively small compared tothe much larger changes in both the enthalpic (Δ[ΔH‡fold] ≈ 11 kcal/mol) and entropic(Δ[T(298K)ΔS‡fold] ≈ 12 kcal/mol) components.

a)3

ln [Keq]

2
1
0
+

K
+
Na

-1
-2
3.20

b)

3.25

3.30
-1

d)

3.40

3

-26
-27
+

-28

c)
c)

3.35

1/T (K ) x 10-25

ln [kfold/]e9

modest (Δ[ΔG‡fold] ≈-1 kcal/mol) decrease in the height of the free energy barrier associated

K Na
3.20

3.25

3.30
-1

+

3.35

1/T (K ) x 10

3.40

3

330
mM

Hfold
kcal/molSfold
kcal/mol/K

K+-20(4)-65(12)

Na+-19(3)-60(8)

330mM

H‡fold

S‡fold

kcal/molkcal/mol/K

K+

2(2)

-60(8)

Na+

0.3(7)-53(2)

Figure 9: (a) van’t
Hoff analysis oftetraloopreceptor
interaction
comparing Na+ andK+ (b) Eyring
analysis of of
tetraloopreceptor
interaction
comparing Na+ andK+ . (c,d) Table ofthermodynamic
parameters fromthe van’t Hoff andEyring analyses

Figure 5.9 (a) van’t Hoff analysis of tetraloop−receptor interaction comparing [Na+] and [K+](b)Eyring analysis of tetraloop−receptor interaction comparing [Na+] and [K+]. (c,d) Table ofthermodynamic parameters from the van’t Hoff and Eyring analyses. As depicted, folding
behavior is largely independent of monovalent cation identity.

122

Indeed, the growth of a strong enthalpic component to the free energy barrier with increasing[Na+] is in surprising opposition to what would be predicted if monovalent-enhanced stability oftertiary interactions were solely the result of a reduction of the entropic penalty associated with
ion uptake or unmixing. Nevertheless, the growth of an endothermic barrier makes entropy thethermodynamic parameter responsible for increased favorability of folding at high monovalentconcentrations. We postulate that these rather dramatic changes in the entropic and enthalpic
components of the free energy barrier may result from monovalent-induced structural changes
within the receptor domain. Specifically, we propose base unstacking and flipping of U19, alongwith the resulting structural changes, as one possible explanation worthy of further exploration.
The results contained within this work support the emerging paradigm that suggests RNA
folding is predominately rate-limited under free energy neutral conditions by a conformational
search (e.g., -TΔS‡fold > ΔH‡fold ≈ 0 kcal/mol) and that addition of monovalent cations reduces theconformational search of RNAs at the potential cost of stabilizing alternative conformations(e.g., -TΔS‡fold ≈ ΔH‡fold > 0 kcal/mol). A thorough examination of other tertiary interactions suchas H-type pseudoknots, larger non-coding RNAs (e.g riboswitches) and perhaps even duplex
formation would be useful to demonstrate the universality of such a paradigm. Additionally, thiswork provides detailed experimental benchmarks for the thermodynamics associated withformation of tertiary interactions in RNA. These benchmarks can be used to test rigorous
theoretical modeling of simple but ubiquitous RNA tertiary interactions in the presence ofmonovalents ions. Finally, this work clearly highlights important directions for future
investigations of the more complex effects associated with mixed-valence cation solutions more
relevant to that of the cellular environment.

123

5.6

Experimental Methods

5.6.1

Experimental Apparatus
Excitation of fluorescent molecules is accomplished by focusing light from a pulsed (20MHz, 10 ps) 532 nm laser to a diffraction-limited spot in the sample with a 1.2 N.A. water
emersion objective. The same objective collects the photons from the sample and directs them toa detection system using a dichroic mirror. A small pinhole rejects fluorescence from outside theconfocal volume thereby reducing background signal. Photons from donor/acceptor fluorescenceare separated by wavelength and polarization using dichroic filters and polarization beam
splitters. The photons are then detected on single-photon avalanche photodiodes (see ref (36) for
details). Photon arrival time information is sent to a time-correlated single-photon counting
module where the fluorescence microtime (i.e. delay with respect to the laser pulse) andmacrotime (time with respect to the start of the experiment) of each incoming photon is recorded.
5.6.2

Temperature Control
Two methods are used to provide temperature control of smFRET samples. The first

involves the use of a microscope stage heater and a heated objective collar to uniformly heat theentire sample (see ref (36) for details). The second involves the use of a newly adapted IR-laser
absorption based heating technique (see ref (66) for details). This technique provides the ability
to control the local temperature of an aqueous sample using focused light from a fiber-coupled
1440 nm continuous-wave laser. While the stage heating method is used for a majority of thedata collection, IR-laser based heating is also explored at all reported values of [Na+]. The twoheating methods used in the present work produce kinetic and thermodynamics results that havepreviously been determined to be indistinguishable from one another(66).

124

5.6.3RNA and Sample Preparation
All nucleic acid sequences for the tetraloop−receptor constructs have been purchasedcommercially. The full construct consists of three strands of oligonucleotides (Figure 5.1): (i) a5) Cy3 labeled RNA strand containing the GAAA tetraloop sequence ii) a 5) Cy5 labeled RNA
strand with a 3) overhang iii) a 5) biotinylated DNA strand complementary to the 3) overhang of
the Cy5 strand. The three strands are annealed and purified using solutions and buffers describedin previously work(10). Thermodynamic and kinetic data was collected using solutions
consisting of: 50 mM hemisodium HEPES, 5.0 mM PCA, 100 nM PCD, 2.0 mM Trolox, 0.1
mM EDTA at pH 7.5 with varying concentrations of NaCl (100-800 mM) to achieve the desired[Na+]. Note the [Na+] comes from added NaCl, buffer, and the oxygen scavenging system—
PCA/PCD.
5.7

Acknowledgment

We would like to thank Drs. Arthur Pardi and Christopher D. Downey for their contributions to
the RNA construct design.

125

5.8

References

1.

Mattick, J. S.; I. V. Makunin. Non-coding RNA, Hum. Mol. Genet., 2006, 15, R17-R29.

2.

Butcher, S. E.; A. M. Pyle. The molecular interactions that stabilize RNA tertiary
structure: RNA motifs, patterns, and networks, Acc. Chem. Res., 2011, 44, 1302-1311.

3.

Ditzler, M. A.; D. Rueda; J. J. Mo; K. Hakansson; N. G. Walter. A rugged free energy
landscape separates multiple functional RNA folds throughout denaturation, Nucl. Acids
Res., 2008, 36, 7088-7099.

4.

Steiner, M.; D. Rueda; R. K. O. Sigel. Ca2+ induces the formation of two distinct
subpopulations of group II intron molecules, Angew. Chem. Int. Ed. Engl. , 2009, 48,
9739-9742.

5.

Xie, Z.; N. Srividya; T. R. Sosnick; T. Pan; N. F. Scherer. Single-molecule studies
highlight conformational heterogeneity in the early folding steps of a large ribozyme,
Proc. Natl. Acad. Sci. USA, 2004, 101, 534-539.

6.

Zhuang, X. W.; H. Kim; M. J. B. Pereira; H. P. Babcock; N. G. Walter; S. Chu.
Correlating structural dynamics and function in single ribozyme molecules, Science,
2002, 296, 1473–1476.

7.

Record, M. T.; W. T. Zhang; C. F. Anderson. Analysis of effects of salts and uncharged
solutes on protein and nucleic acid equilibria and processes: A practical guide torecognizing and interpreting polyelectrolyte effects, Hofmeister effects, and osmotic
effects of salts, Adv. Protein. Chem., 1998, 51, 281-353.

8.

Privalov, P. L.; O. B. Ptitsyn; Birshtei.Tm. Determination of stability of DNA double
helix in an aqueous medium, Biopolymers, 1969, 8, 559-571.

9.

Manning, G. S. Application of polyelectrolyte limiting laws to helix-coil transition of
DNA .1. Excess univalent cations, Biopolymers, 1972, 11, 937-947.

10.

Downey, C. D.; J. L. Fiore; C. D. Stoddard; J. H. Hodak; D. J. Nesbitt; A. Pardi. Metal
ion dependence, thermodynamics, and kinetics for intramolecular docking of a GAAA
tetraloop and receptor connected by a flexible linker, Biochemistry, 2006, 45, 3664–3673.

11.

Heilman-Miller, S. L.; J. Pan; D. Thirumalai; S. A. Woodson. Role of counterion
condensation in folding of the Tetrahymena ribozyme II. Counterion-dependence offolding kinetics, J. Mol. Biol., 2001, 309, 57–68.

12.

Heilman-Miller, S. L.; D. Thirumalai; S. A. Woodson. Role of counterion condensation
in folding of the Tetrahymena ribozyme. I. Equilibrium stabilization by cations, J. Mol.
Biol., 2001, 306, 1157–1166.

13.

Nixon, P. L.; D. P. Giedroc. Energetics of a strongly pH dependent RNA tertiary
structure in a frameshifting pseudoknot, J. Mol. Biol., 2000, 296, 659–671.

126

14.

Takamoto, K.; Q. He; S. Morris; M. R. Chance; M. Brenowitz. Monovalent cations
mediate formation of native tertiary structure of the Tetrahymena thermophila ribozyme,
Nat. Struct. Biol., 2002, 9, 928-933.

15.

Uchida, T.; K. Takamoto; Q. He; M. R. Chance; M. Brenowitz. Multiple monovalent iondependent pathways for the folding of the L-21 Tetrahymena thermophila ribozyme, J.
Mol. Biol., 2003, 328, 463-478.

16.

Ke, A.; F. Ding; J. D. Batchelor; J. A. Doudna. Structural roles of monovalent cations in
the HDV ribozyme, Structure, 2007, 15, 281-287.

17.

Jiang, Y. F.; M. Xiao; P. Yin; Y. Zhang. Monovalent cations use multiple mechanisms toresolve ribozyme misfolding, RNA, 2006, 12, 561-566.

18.

Lambert, D.; D. Leipply; R. Shiman; D. E. Draper. The influence of monovalent cation
size on the stability of RNA tertiary structures, J. Mol. Biol., 2009, 390, 791-804.

19.

Shiman, R.; D. E. Draper. Stabilization of RNA tertiary structure by monovalent cations,J. Mol. Biol., 2000, 302, 79–91.

20.

Uchida, T.; Q. He; C. Y. Ralston; M. Brenowitz; M. R. Chance. Linkage of monovalentand divalent ion binding in the folding of the P4-P6 domain of the Tetrahymena
ribozyme, Biochemistry, 2002, 41, 5799-5806.

21.

Basu, S.; R. P. Rambo; J. Strauss-Soukup; J. H. Cate; A. R. Ferre-D'Amare; S. A.
Strobel; J. A. Doudna. A specific monovalent metal ion integral to the AA platform of the
RNA tetraloop receptor, Nat. Struct. Biol., 1998, 5, 986–992.

22.

Wang, Y. X.; M. Lu; D. E. Draper. Specific ammonium ion requirement for functional
ribosomal-RNA tertiary structure, Biochemistry, 1993, 32, 12279-12282.

23.

Draper, D. E. RNA folding: Thermodynamic and molecular descriptions of the roles ofions, Biophys. J., 2008, 95, 5489-5495.

24.

Anderson, C. F.; M. T. Record. Salt nucleic-acid interactions, Annu. Rev. Phys. Chem.,
1995, 46, 657–700.

25.

Tinoco, I.; C. Bustamante. How RNA folds, J. Mol. Biol., 1999, 293, 271–281.

26.

Sosnick, T. R.; T. Pan. RNA folding: Models and perspectives, Curr. Opin. Struct. Biol.,
2003, 13, 309–316.

27.

Woese, C. R.; S. Winker; R. R. Gutell. Architecture of ribosomal-RNA: "Constraints on
the sequence of Tetraloops", Proc. Natl. Acad. Sci. USA, 1990, 87, 8467-8471.

28.

Hodak, J. H.; C. D. Downey; J. L. Fiore; A. Pardi; D. J. Nesbitt. Docking kinetics andequilibrium of a GAAA tetraloop-receptor motif probed by single-molecule FRET, Proc.
Natl. Acad. Sci. USA, 2005, 102, 10505–10510.

127

29.

Cate, J. H.; A. R. Gooding; E. Podell; K. H. Zhou; B. L. Golden; C. E. Kundrot; T. R.
Cech; J. A. Doudna. Crystal structure of a group I ribozyme domain: Principles of RNA
packing, Science, 1996, 273, 1678–1685.

30.

Batey, R. T.; R. P. Rambo; J. A. Doudna. Tertiary motifs in RNA structure and folding,Angew. Chem. Int. Ed., 1999, 38, 2327–2343.

31.

Costa, M.; F. Michel. Frequent use of the same tertiary motif by self-folding RNAs,
Embo. J., 1995, 14, 1276–1285.

32.

Vander Meulen, K. A.; J. H. Davis; T. R. Foster; T. Record; S. E. Butcher.
Thermodynamics and folding pathway of tetraloop receptor-mediated RNA helicalpacking, J. Mol. Biol., 2008, 384, 702-717.

33.

Young, B. T.; S. K. Silverman. The GAAA tetraloop-receptor interaction contributes
differentially to folding thermodynamics and kinetics for the P4-P6 RNA domain,
Biochemistry, 2002, 41, 12271–12276.

34.

Sattin, B. D.; W. Zhao; K. Travers; S. Chut; D. Herschlag. Direct measurement of tertiary
contact cooperativity in RNA folding, J. Am. Chem. Soc., 2008, 130, 6085–6087.

35.

Fiore, J. L.; J. H. Hodak; O. Piestert; C. D. Downey; D. J. Nesbitt. Monovalent andDivalent Promoted GAAA-Tetraloop–Receptor Tertiary Interactions from Freely
Diffusing Single-Molecule Studies, Biophys. J., 2008, 95, 3892-3905.

36.

Fiore, J. L.; B. Kraemer; F. Koberling; R. Erdmann; D. J. Nesbitt. Enthalpy-Driven RNA
Folding: Single-Molecule Thermodynamics of Tetraloop–Receptor Tertiary Interaction,
Biochemistry, 2009, 48, 2550–2558.

37.

Butcher, S. E.; T. Dieckmann; J. Feigon. Solution structure of a GAAA tetraloop receptor
RNA, Embo. J., 1997, 16, 7490–7499.

38.

Davis, J. H.; M. Tonelli; L. G. Scott; L. Jaeger; J. R. Williamson; S. E. Butcher. RNA
helical packing in solution: NMR structure of a 30 kDa GAAA tetraloop-receptor
complex J. Mol. Biol., 2005, 351, 371–382.

39.

Heus, H. A.; A. Pardi. Structural features that give rise to the unusual stability of RNA
hairpins containing GNRA loops, Science, 1991, 253, 191-194.

40.

Jucker, F. M.; H. A. Heus; P. F. Yip; E. H. M. Moors; A. Pardi. A network ofheterogeneous hydrogen bonds in GNRA tetraloops, J. Mol. Biol., 1996, 264, 968-980.

41.

Aleman, E. A.; R. Lamichhane; D. Rueda. Exploring RNA Folding One Molecule at aTime, Curr. Opin. Chem. Biol., 2008, 12, 647-654.

42.

Bokinsky, G.; X. W. Zhuang. Single-molecule RNA folding, Acc. Chem. Res., 2005, 38,
566–573.

128

43.

Blanco, M.; N. G. Walter. Analysis of complex single-molecule FRET time trajectories,
Methods Enzymol., 2010, 472, 153-178.

44.

Bartley, L. E.; X. W. Zhuang; R. Das; S. Chu; D. Herschlag. Exploration of theTransition State for Tertiary Structure Formation Between an RNA Helix and a Large
Structured RNA, J. Mol. Biol., 2003, 328, 1011–1026.

45.

Zhou, Y. J.; X. W. Zhuang. Robust reconstruction of the rate constant distribution using
the phase function method, Biophys. J., 2006, 91, 4045-4053.

46.

Leipply, D.; D. Lambert; D. E. Draper. Ion-RNA interactions: Thermodynamic analysisof the effects of mono- and divalent ions on RNA conformational equilibria, Methods
Enzymol., 2009, 469, 433-463.

47.

Bokinsky, G.; D. Rueda; V. K. Misra; M. M. Rhodes; A. Gordus; H. P. Babcock; N. G.
Walter; X. W. Zhuang. Single-molecule transition-state analysis of RNA folding, Proc.
Natl. Acad. Sci. USA, 2003, 100, 9302–9307.

48.

Fiore, J. L.; E. D. Holmstrom; D. J. Nesbitt. An Entropic origin of Mg2+-Facilitated RNA
Folding, Proc. Natl. Acad. Sci. USA, 2011, (Accepted).

49.

Hanggi, P.; P. Talkner; M. Borkovec. Reaction-rate theory - 50 years after Kramers, Rev.
Mod. Phys., 1990, 62, 251-341.

50.

Vander Meulen, K. A.; S. E. Butcher. Characterization of the kinetic and thermodynamic
landscape of RNA folding using a novel application of isothermal titration calorimetry,
2011, DOI:10.1093/nar/gkr894.

51.

Davis, J. H.; T. R. Foster; M. Tonelli; S. E. Butcher. Role of metal ions in the tetraloopreceptor complex as analyzed by NMR, RNA, 2007, 13, 76-86.

52.

Buchmueller, K. L.; A. E. Webb; D. A. Richardson; K. M. Weeks. A collapsed nonnative RNA folding state, Nat. Struct. Biol., 2000, 7, 362-366.

53.

Swisher, J. F.; L. H. J. Su; M. Brenowitz; V. E. Anderson; A. M. Pyle. Productive folding
to the native state by a group II intron ribozyme, J. Mol. Biol., 2002, 315, 297-310.

54.

Silverman, S. K.; T. R. Cech. An early transition state for folding of the P4-P6 RNA
domain, RNA, 2001, 7, 161–166.

55.

Pljevaljcic, G.; D. Klostermeier; D. P. Millar. The tertiary structure of the hairpin
ribozyme is formed through a slow conformational search, Biochemistry, 2005, 44, 48704876.

56.

Szewczak, A. A.; E. R. Podell; P. C. Bevilacqua; T. R. Cech. Thermodynamic stability of
the P4-P6 domain RNA tertiary structure measured by temperature gradient gelelectrophoresis, Biochemistry, 1998, 37, 11162–11170.

129

57.

Vander Meulen, K. A.; S. E. Butcher. Characterization of the kinetic and thermodynamic
landscape of RNA folding using a novel application of isothermal titration calorimetry,Nucl. Acids Res., 2012, 40, 2140-2151.

58.

Holbrook, J. A.; M. W. Capp; R. M. Saecker; M. T. Record. Enthalpy and heat capacitychanges for formation of an oligomeric DNA duplex: Interpretation in terms of coupledprocesses of formation and association of single-stranded helices, Biochemistry, 1999, 38,
8409-8422.

59.

Takach, J. C.; P. J. Mikulecky; A. L. Feig. Salt-dependent heat capacity changes for RNA
duplex formation, J. Am. Chem. Soc., 2004, 126, 6530-6531.

60.

Qin, P. Z.; J. Feigon; W. L. Hubbell. Site-directed spin labeling studies reveal solution
conformational changes in a GAAA tetraloop receptor upon Mg2+-dependent docking ofa GAAA tetraloop, J. Mol. Biol., 2005, 351, 1-8.

61.

Lambert, D.; D. Leipply; D. E. Draper. The osmolyte TMAO stabilizes native RNA
tertiary structures in the absence of Mg2+: Evidence for a large barrier to folding from
phosphate dehydration, J. Mol. Biol., 2010, 404, 138-157.

62.

Breslauer, K. J.; J. M. Sturtevant. Calorimetric investigation of single stranded basestacking in ribo-oligonucleotide-A7, Biophys. Chem., 1977, 7, 205-209.

63.

Seol, Y.; G. M. Skinner; K. Visscher. Elastic properties of a single-stranded charged
homopolymeric ribonucleotide, Phys. Rev. Lett., 2004, 93, 118102.

64.

Michelson, A. M.; C. Monny. Polynucleotides .8. Base stacking in polyuridylic acid,
Proc. Natl. Acad. Sci. U.S.A., 1966, 56, 1528-1534.

65.

Fiore, J. L.; E. D. Holmstrom; L. R. Feigland; J. H. Hodak; D. J. Nesbitt. The Role ofCounterion Valence and Size in the GAAA Tetraloop-Receptor Docking/Undocking
Kinetics, 2011, (In preperation).

66.

Holmstrom, E. D.; D. J. Nesbitt. Real-time infrared overtone laser control of temperaturein picoliter H2O samples: "Nanobathtubs" for single-molecule microscopy, J. Phys.
Chem. Lett., 2010, 1, 2264–2268.

130

Chapter 6

Kinetic and Thermodynamic Origins of Osmolyte-Influenced Nucleic Acid Folding*

6.1

Introduction
The folding free energy for nucleic acid is significantly influenced by the presence ofcosolutes. Some of the most prominent cosolutes are metal ions, which are known to beimportant for a large number of nucleic acid conformational transitions ranging in complexity
from short, single-strand oligonucleotides to large, highly organized RNAs like the large
ribosomal subunit(1,2). However, metal ions are not the only cosolutes that can alter theenergetics of nucleic acid folding. Many highly soluble organic small molecules—known as
osmolytes—can also affect the stability of various nucleic acid structural motifs(3).
Early work on the biological importance of osmolytes has demonstrated that they help
living cells respond to changes in osmotic pressure(4,5). However, numerous experiments haveshown that high concentrations of osmolytes can either facilitate or inhibit folding of nucleic
acids and proteins, depending on the chemical nature of the osmolyte. As intracellularconcentrations change in response to osmotic stress, the effects of these cosolutes on theenergetics of structured proteins and nucleic acids must therefore be counterbalanced in order tomaintain homeostasis. Previous biological studies have validated this idea by demonstrating thatsome organisms are able to adjust intracellular concentrations of stabilizing osmolytes to offset
the effect of urea(6,7).

*

To be submitted131

Because osmolytes are known to alter the stability of biomolecules, a large number ofscientific investigations have centered on studying their ability to influence protein (8-11) andnucleic acid (12-17) conformational transitions. Urea is one of the most prominent osmolytes andtends to destabilize folded proteins(18-20) and nucleic acids(21-23), where as trimethylamine Noxide (TMAO) is an osmolyte that has been shown to stabilize these biomolecules(13,22,24,25).
Recently, the interactions between osmolytes and nucleic acids have begun to attract the same
degree of attention as their protein-based analogues. To follow suit, this work reports on the first,
to our knowledge, detailed kinetic investigation of the topic using single-molecule fluorescence
techniques.
Over the years, a number of experimental and theoretical efforts regarding TMAO(13,26)
and urea(14,27) have brought about a more physical understanding of such nucleic acidosmolyte interactions. Inspired by quantitative thermodynamic work on proteins(11), recent bulk
ensemble studies indicate that osmolytes can interact with any of the three components of a
nucleic acid—base, sugar, and phosphate(12-14). Specifically, these ensemble experiments havedemonstrated that urea destabilizes RNA folding transitions by forming favorable interactions
with all three constituents, which increase in strength from urea-phosphate to ureanucleobase(14). Because the unfolded conformations of most nucleic acids tend to have moresolvent accessible surface area (SASA), the presence of urea will therefore tend to preferentiallystabilize such unfolded conformations relative to those that are folded and more compact (Figure6.1). This results in a net increase in the free energy change (ΔG°fold) between the unfolded andfolded conformation, which is driven primarily by SASA-mediated burial of nucleobases. For
this reason, urea is referred to as a destabilizing osmolyte. Conversely, the presence of theosmolyte TMAO gives rise to precisely the opposite effect, specifically a net decrease in ΔG°fold.

132

This results primarily from unfavorable interactions between the osmolyte and the various
surfaces of a nucleic acid, which range in strength from TMAO- phosphate to the rather weak
TMAO-sugar interactions(13). As described above, nucleic acids tend to become more compact
when they fold and, as a result, phosphate SASA is often buried. Therefore, the presence of
TMAO yields a preferential destabilization of the unfolded species, which results in a net
stabilization of the folding transition (Figure6.1). Hence, TMAO is characterized as a stabilizing
osmolyte.

Figure 6.1 Free energy diagram representing the effect of TMAO or urea on the stability of theunfolded (U) or folded (F) structures of a nucleic acid. TMAO has a more unfavorable
interaction with the unfolded structure than the folded structure, which results in a net decrease
in ΔG°fold. Urea has a more favorable interaction with the unfolded conformation than folded
conformation, which gives rise to a net increase in ΔG°fold.
To date, most experimental studies that investigate the influence of osmolytes on
structured nucleic acid have been based on ensemble techniques (e.g., calorimetry or UVspectroscopy) that only provide information about the equilibrium constant between folded andunfolded species (e.g., Keq = [folded]/[unfolded]=). Although these techniques have been
extremely informative, they lack the ability to partition the observed equilibrium effects into
components related to the folding/unfolding kinetics (e.g., Keq = kfold/kunfold). Because of this, the133kinetic origin of osmolyte-influenced nucleic acid folding is not well known and is, therefore, themajor focus of this study.
Single-molecule methods(28) are ideally suited to address these questions by providing
kinetic information about the folding and unfolding rate constants of various nucleic acidconformational transitions. To this end, single-molecule fluorescence resonance energy transfer(29-32) (smFRET) is used to determine the kinetic origins of TMAO-facilitated and ureainhibited nucleic acid folding. Specifically, two fluorescently labeled constructs (Figure 6.2)
serve as models systems for both types of nucleic acid folding transitions—secondary (2°)
structure formation (e.g., duplex association/dissociation) and tertiary (3°) structure formation(e.g., loop docking/undocking). The first model construct is a previously characterized(33) 8 bp
DNA duplex that is used to elucidate the role of osmolytes in the formation of nucleic acid 2°
structures (Figure 6.2a,b). The second is an RNA construct that isolates the GAAA tetraloop—
tetraloop receptor (TL-TLR) interaction from the Tetrahymena ribozyme. This ubiquitous loopbulge interaction has undergone extensive biophysical characterization at the single-molecule
level (34-40) and is used to study the influence of osmolytes on RNA 3° structure formation(Figure 6.2b,c). Using smFRET techniques to study these model systems, we show that the
kinetic origin of TMAO-facilitated nucleic acid folding arises from an increase in kfold and acorresponding decrease in kunfold for both 2° and 3° structure formation. The precise opposite istrue for the kinetic origin of urea-inhibited nucleic acid structure formation, which results from adecrease in kfold and an increase in kunfold, regardless of the nucleic acid construct.
6.2

Results
The two nucleic acid constructs are chosen to serve as model systems for a variety ofreasons. First and foremost, they broadly represent the two types of structure formation in134

nucleic acids—secondary (2°) and tertiary (3°). Specifically, canonical Watson-Crickinteractions within a double helix are one of the defining characteristics of nucleic acid 2°
structures.

Figure 6.2 Cartoon representation of the two smFRET nucleic acid constructs used throughout
the study. The DNA duplex construct (a and b) is intended to be a model system for nucleic acidsecondary structure, where as the TL-TLR construct (c and d) serves as a model for tertiary
interactions in structured RNAs. For both constructs, formation of the interaction (b and d),referred to as folding, results in an increase in EFRET(a and c).
Alternatively, the interaction between the GAAA tetraloop and the tetraloop receptor (TL-TLR)
contains both non-canonical hydrogen bonding and base stacking between a loop and an internalbulge, which are among some of the distinguishing features of 3° structures. These tertiary
contacts frequently orient and stabilize the global architecture and three-dimensional
conformation of large structured nucleic acids. An additional reason for using these two

135

constructs as model systems is that both the 8 bp DNA duplex(33) and the TL-TLR have beenpreviously characterized using single-molecule techniques(34-40), which establishes a wellunderstood baseline for the folding kinetics in the absence of osmolytes. Lastly, both interactions
are capable of forming in the absence of Mg2+. This allows the equilibrium constant for the two
conformational transitions to easily be adjusted via the [KCl] and also limits the number ofcosolutes in solution, therefore reducing the potential for complex cooperative effects observedin more complex solutions.
6.2.1

Folding in the Absence of Osmolytes
Individual time-trajectories for surface immobilized TL-TLR constructs demonstrate two-state behavior (Figure 6.3a) with well-defined high and low FRET states (e.g., docked !
undocked). The 8bp DNA duplex exists in a bimolecular equilibrium (e.g., strand 3+ dsDNA
scaffold ! duplex) with a concentration-dependent rate constant for duplex association. To
facilitate determination of this rate constant, experiments are performed at a fixed concentration(200 nM) of the freely diffusing Cy5, as previously reported(33). Surface immobilized
trajectories of the 8bp duplex also exhibit two-state behavior (Figure 6.3b). The well-separated
FRET states for each of the two constructs make it easy to determine dwell times using apreviously described thresholding routine(39). The experimentally measured dwell times areanalyzed via a cumulative distribution function(41-43) to determine the rate constants for eachconstruct, which establishes a point-of-comparison for experiments that are conducted using
solutions containing high concentrations of osmolytes. For ease of discussion, the forward rate
constant for both systems (i.e. duplex association and tetraloop docking) will be referred to as thefolding rate constant (kfold), whereas the reverse rate constant will be referred to as the unfolding
rate constant (kunfold).

136

Figure 6.3 Representative donor (green) and acceptor (red) fluorescence trajectories for (a) the
TL-TLR and (b) the DNA duplex. Fluorescence data is use to calculate the FRET trajectories(black), which are then analyzed to measure the folding (kfold) and unfolding (kunfold) rate
constants for each of the nucleic acid constructs.
To further facilitate kinetic and thermodynamic comparisons between constructs, KClconcentrations are adjusted so that ΔG° in the absence of any osmolytes is ≈ 0 for both folding
transitions (i.e., 50 mM and 100 mM for the TL-TLR and 8 bp DNA duplex, respectively). The
following four sections highlight the important kinetic and thermodynamic observations
associated with osmolyte-influenced nucleic acid folding.
6.2.2

Osmolyte-Dependent Folding Kinetics
Titrations of both TMAO and urea clearly demonstrate the kinetic origin of osmolyte-

influenced nucleic acid folding (Figure 6.4). The presence of TMAO increases kfold (Figure 6.4a)and decreases kunfold (Figure 6.4b) for both nucleic acid constructs. Together, these kinetic effects
contribute to an increase in Keq and more favorable folding.

137

Figure 6.4 Kinetic origin of osmolyte- influenced nucleic acid folding. The stabilizing osmolyte
TMAO (blue box) increases the (a) folding rate constant (kfold, red) and decreases the (b)unfolding rate constant (kunfold, green) for both the GAAA tetraloop—receptor (triangle) andDNA duplex (circle) constructs. The destabilizing osmolyte, urea, decreases the (c) folding rate
constant (kfold, red) and increases the (d) unfolding rate constant (kunfold, green) for bothconstructs.
The exact opposite effect results from the presence of urea, where elevated concentrations
decrease kfold (Figure 6.4c) and increase kunfold (Figure 6.4d) for both the 8 bp DNA duplex and
the TL-TLR. Interestingly, the qualitative trends highlighted above are quantitatively different.
These distinctions are exemplified by the following two observations: (i) TMAO accelerates kfold
for the TL-TLR more effectively that it does for the 8 bp DNA duplex (Figure 6.4a, red triangles
vs circles) and (ii) urea decelerates kunfold for the 8 bp DNA duplex more effectively that it does
for the TL-TLR (Figure 6.4d, green triangles vs circles).

1386.2.3Osmolyte-Dependent Folding Free Energies (m-value analysis)With knowledge of both rate constants, the free energy change for folding (ΔG°fold) canreadily be calculated from the equilibrium constant (Keq = kfold/kunfold) using the Gibbs equation(ΔG° = RTln[Keq]). As empirically observed for nucleic acids(21,44,45) and proteins(46,47),
the folding free energy change is linearly-dependent on osmolyte concentration (Figure 6.5).
This dependence (∂ΔG°/∂[osmolyte]) is often characterized by the m-value, which serves as ameasure of the folding transition’s sensitivity to the presence of an osmolyte. From these mvalues, it is apparent that: (i) TMAO is much more effective at stabilizing the TL-TLR than the 8bp DNA duplex and (ii) urea destabilizes both model constructs by an equal amount (Figure 6.5).

Figure 6.5 Osmolyte-dependent folding free energy change for (a) the DNA duplex and (b) the
TL-TLR constructs. For both constructs, addition of TMAO (blue stars) always stabilizes thefolding transitions, where as addition of urea (orange squares) destabilizes the folding transitions
as indicated by the dependence of ΔG° on the osmolyte concentration.
Both of these observations are in complete accordance with previous equilibrium-based
ensemble studies, which reveal that urea can effectively destabilize both 2° and 3° interactions,139

where as TMAO primarily stabilizes 3° structure formation. Additionally, this experimental
agreement further validates the use of single-molecule experiments to study osmolyte-influenced
nucleic acid conformational transitions with the intent to partition equilibrium effects into theirkinetic components.
6.2.4

Osmolyte-Dependent Thermodynamics for the 8 bp DNA Duplex
To more rigorously understand the kinetic origin of osmolyte-influenced folding of

nucleic acids, temperature-dependent single-molecule experiments have been performed to parsethe free energy change into its enthalpic and entropic components (ΔG° = ΔH° TΔS°)(40,48).
Briefly, the inverse temperature-dependence of ln[Keq] can be used to provide information about
ΔH° and ΔS° for the two model systems in the form of a van’t Hoff analysis. Similarly, theinverse temperature-dependence of ln[k] provides information about the enthalpic and entropic
components of the free energy barrier (ΔH‡ and ΔS‡) in what is referred to as an Eyring analysis.
In a plot of ln[Keq] vs 1/T (Figure 6.6), the slope represents −ΔH°/R and the intercept,
corresponds to ΔS°/R, where R is the Gas constant (≈ 1.987 cal mol-1 K-1). For the 8 bp DNA
duplex, neither of the 1M osmolyte solutions substantially changes ΔH° (Figure 6.6a). This
indicates that the thermodynamic origin of TMAO- and urea-influenced 2° structure formation is
predominately entropic. Furthermore, the Eyring analyses demonstrate that there is a much largerchange in ΔS‡unfold than ΔS‡fold, which implies that this entropic origin is associated with theunfolding process (Supplemental Figure S6.1b). Explicitly stated, TMAO stabilizes the 8 bp
DNA duplex by making the unfolding barrier less entropically rewarding. Conversely, urea
destabilizes this conformational transition by making the unfolding barrier more entropicallyrewarding. As a testament to the validity of these single-molecule experiments, thethermodynamics results for the 8 bp DNA duplex are in excellent agreement with the140corresponding kinetic data, which demonstrates that both osmolytes more prominently affectkunfold rather than kfold (Figure 6.4, circles).

Figure 6.6 van’t Hoff plots for the osmolyte- influenced folding of both (a) the DNA duplex and(b) the TL-TLR interaction. Solid lines in (a) correspond to a global fit with a common slope, as
discussed in the text, to reduce parameter correlation between the three data sets: 1 M TMAO
(blue stars), 0 M osmolyte (pink circles), and 1 M urea (orange squares). The solid lines for the
TL-TLR interaction (b) are also the result of a global fit of the 0 M osmolyte and 1 M urea data
sets in order to reduce parameter correlation, while the solid line for the 1M TMAO data set
represents an independent fit of the data. See Table 6.1 for values of the fitted parameters.

141

Table 6.1 Values for the van’t Hoff analysis of the osmolyte-influenced folding nucleic acids.
Additional information about the methods of data analysis for the van’t Hoff plots can be foundin Section 6.3.4. See Figure 6.6 for experimental data.

Supplemental Figure S6.1 Temperature-dependent kinetic data for the (a) folding and (b)unfolding rate constants for the DNA duplex. The data are analyzed using transition state theory.
Solid lines in correspond to a global fit with a common slope, as discussed in the text, to reduceparameter correlation. The large entropic offset in the temperature-dependence of the unfolding
rate constants (b) suggests that most of the change in ΔS° arises from changes in the entropic
barrier to unfolding (ΔS‡unfold). The data table (c) contains values for the transition state theory
analysis of the temperature-dependent kinetic data for the DNA duplex. Fitting methods are
identical to those used in the van’t Hoff analysis (Figure 6.6), which is discussed in Section
6.3.4.

142

6.2.5Osmolyte-Dependent Thermodynamics for the TL-TLR
The van’t Hoff and Eyring analyses of the TL-TLR highlight key similarities anddifferences between the two osmolytes. Additionally, when compared to results of thethermodynamic studies of the 8 bp DNA duplex, these observations also highlight similarities
and differences between the two types of conformational transitions. As was the case for theurea-inhibited folding of the 8 bp DNA duplex, the presence of 1 M urea has little affect on the
ΔH° for the TL-TLR interaction (Figure 6.6b), which, again, indicates that entropy is thethermodynamic parameter responsible for destabilization. However, unlike the 8bp DNA duplex,
most of this change in ΔS° arises from the folding process (Supplemental Figure S6.2). The ureadependent thermodynamics for the TL-TLR construct are remarkably well supported by thecorresponding kinetics, which demonstrate that destabilization occurs primarily via a reductionof kfold (Figure 6.4c, triangles). Together, the kinetic and thermodynamic results from the TLTLR experiments imply that urea destabilizes this interaction by increasing the entropic costassociated with the folding free energy barrier.
Interestingly, elevated concentrations of TMAO change both the ΔH° (slope) and the ΔS°(intercept) associated with formation of the TL-TLR interaction (Figure 6.6b), which is also
demonstrated in Eyring analyses (Supplemental Figure S6.2a). This is surprising because TMAO
does not affect the ΔH° for the 8bp DNA duplex. The prominence of both an enthalpic andentropic component to TMAO-facilitated formation of the TL-TLR suggests that this osmolyte
interacts differently with the two nucleic acid constructs, presumably as a result of considerably
different changes in the phosphate SASA.

143

Supplemental Figure S6.2 Temperature- dependent kinetic data for the (a) folding and (b)unfolding rate constants for the TL-TLR. The data are analyzed using transition state theory.
Solid lines for 1 M urea (orange squares) and 0 M osmolyte (pink circles) correspond to a global
fit with a common slope, as discussed in the text, to reduce parameter correlation. The large
entropic offset for 1 M urea in the temperature-dependence of the folding rate constant (a)suggests that most of the change in ΔS° arises from changes in the entropic barrier to folding(ΔS‡fold). The distinct thermodynamic behavior of TMAO- facilitated formation of the TL-TLR
interaction is discussed in the text. The data table (c) contains values for the transition state
theory analysis of the temperature-dependent kinetic data for the DNA duplex. Fitting methods
are identical to those used in the van’t Hoff analysis (Figure 6.6), which is discussed in Section
6.3.4.
6.3

Discussion
Ensemble thermal denaturation experiments have demonstrated that the m-value, which isa measure of the sensitivity of ΔG° to osmolyte concentration, is correlated with the differencesin solvent accessible surface area (SASA) between the folded and unfolded structures.
Specifically, the m-values for TMAO and urea are primarily mediated by changes in phosphateand nucleobase SASA, respectively. Vapor pressure osmometry (VPO) experiments haverevealed that TMAO primarily forms unfavorable interactions with the phosphate backbone, and144

to a lesser extent, the nucleobases of a nucleic acid(13). This is different from the favorable
interaction between urea and the nucleobase of an oligonucleotide(14). Accordingly, TMAO will
preferentially destabilize conformations with more phosphate SASA, whereas urea will betterstabilize structures with greater base SASA (Figure 6.1). Unfolded conformations commonly
have more phosphate and base SASA than their folded counterparts, which means that TMAO
will stabilize nucleic acid folding transitions, whereas urea will destabilize them (Figure 6.1).
6.3.1 TMAO-Dependent Folding Free Energies
The folding/unfolding rate constants (Figure 6.4) are used to calculate equilibrium
constants (Keq = kfold/kunfold), which are used to determine the osmolyte-dependence of the freeenergy change for folding (m-value). These results clearly indicate that TMAO is better able tostabilize 3° (e.g., TL-TLR), rather than 2° (e.g., DNA duplex), structural transitions (Figure 6.5).
Previous calculations based on structural models of duplex nucleic acids predict very littlechange in phosphate SASA upon formation of a double helix(12), which would render TMAO an
ineffective stabilizing osmolyte. However, VPO experiments(13) indicate that there are someweak, but still energetically unfavorable, interactions between TMAO and nucleobases thatwould result in a net stabilization of the duplex based on the change in nucleobase SASA.
Conversely, the substantial change in phosphate SASA (≈ 67 Å2) for the TL-TLR receptor
interaction (13) clearly explains why TMAO is more effective at stabilizing this structuraltransition. These results bolster the notion that, when changes in phosphate SASA are
substantial, TMAO can efficiently stabilize conformational transitions. However, when thesechanges are miniscule, TMAO can still facilitate folding, although less efficiently, based on
differences in nucleobase SASA.

145

6.3.2 Urea-Dependent Folding Free Energies
Surprisingly, the m-values for the TL-TLR and 8 bp DNA duplex interactions are very
similar (Figure 6.5), which suggests nearly identical changes in nucleobase SASA for the two
folding transitions. Additionally, previous ensemble experiments on similar constructs undercomparable conditions yield qualitatively consistent m-values for urea-inhibited folding. A10 bp
RNA duplex at 0.5 M NaCl has an m-value = 0.74(4) kcal mol-1 molar-1(21) and a bimolecular,
dimeric TL-TLR construct has an extrapolated m-value ≈ 0.64 kcal mol-molal-1 at 85 mM KCl(14). Both of these previously measured values are within the experimental uncertainty of thesingle-molecule measurements reported in this study. The excellent agreement between m-values
measured using single-molecule techniques and UV-melting experiments further supports thenotion that urea is a strongly destabilizing osmolyte for both 2° and 3° folding transitions, which
results primarily from favorable interactions with the nucleobase SASA of nucleic acids. Despite
the similar m-values, which are related to the folding equilibria, the pronounced structuraldifferences between the TL-TLR and the 8bp DNA duplex make the urea-dependence of the rate
constants very different.
6.3.3

Kinetic Origins of Osmolyte-Influenced Nucleic Acid FoldingThis work details the first, to our knowledge, demonstration of the kinetic origins

associated with osmolyte-influenced nucleic acid folding. The observed increase in stability offolded nucleic acid structures at elevated concentrations of TMAO is the result of an increase in
kfold (Figure 6.4a) and a decrease in kunfold (Figure 6.4b). Alternatively, the decrease in stability
associated with addition of urea occurs because of a decrease in kfold (Figure 6.4c) and anincrease in kunfold (Figure 6.4d).

146

The ideology of preferential interactions with base, sugar, and phosphate SASA of
nucleic acids can also be applied to osmolyte-dependence of the rate constants. Such an analysis
provides information about changes in SASA that result from forming the transition state. For
example, the observation that kfold for the 8 bp DNA duplex construct is only slightly increasedby the presence of TMAO (Figure 6.4a, circles) indicates that there is very little reduction inphosphate SASA upon reaching the transition state. Conversely, the kfold for TL-TLR constructstrongly accelerated by TMAO (Figure 6.4a, triangles), which implies that there is aconsiderable reduction in phosphate SASA that occurs while crossing the folding free energy
barrier.
Similarly, the urea-dependence of the rate constants can be used to make inferences about
where along the reaction coordinate the burial of nucleobase SASA occurs for the two different
conformational transitions. Because kfold for the TL-TLR interaction is much more sensitive tourea than kunfold (Figures 6.4c, 6.5d) it evident that most of the change in nucleobase SASA
occurs prior to formation of the transition state. This is in contrast with previous smFRET studies
of urea-induced denaturation of other RNA folding motifs (i.e., the human telomerase RNA
pseudoknot(49), the adenine riboswitch(50), and the P4-P6 domain of the Tetrahymena
ribozyme(42)) where addition of urea predominantly affects the unfolding rate constant. It isinteresting to note that the GAAA tetraloop—tetraloop receptor is one of two tertiary interactionsin the P4-P6 domain and that the inconsistencies between these two RNA constructs are likely
the result of vastly different folding reaction coordinates, specifically regarding changes innucleobase SASA.
Not only do these results demonstrate the kinetic origins of osmolyte-influenced nucleic
acid folding (Figure 6.4), but also that osmolytes, specifically TMAO and urea, can be147experimental tools capable of providing low-resolution information about changes in SASA
across the entire folding landscape. Accordingly, the large amount of nucleobase and phosphate
SASA buried upon formation of the transition state for the TL-TLR interaction is evidence for atransition state where the GAAA tetraloop and the receptor are proximal to one another. Such astructural depiction is certainly consistent with independent single-molecule experiments that
propose a compact transition state for the GAAA tetraloop—receptor interaction(40).
6.3.4

Osmolyte-Dependent Entropy Changes for the 8bp DNA Duplex
Temperature-dependent smFRET experiments have been preformed to obtain insights

into the enthalpic and entropic components (i.e., thermodynamics origin) of osmolytesinfluenced nucleic acid folding. As indicated by the vertically-offset, parallel data sets in the
van’t Hoff plots for the 8 bp DNA duplex (Figure 6.6a), neither 1 M TMAO nor 1 M urea
significantly alter the folding enthalpy. This observation demonstrates that ΔH° is insensitive tothe presence of osmolytes (Table 6.1) and that the predominant source of the osmolytedependent ΔG° (m-value) results from entropic considerations. Previous calorimetricexperiments that examined the effect of destabilizing osmolytes (e.g., urea) on DNA melting
have similarly concluded that entropic terms dominate, giving rise to the observed decrease induplex stability (15). Additionally, computational simulations of an RNA hairpin (e.g., 2°
structure) in the presence of TMAO suggest that, if the interaction between the two isunfavorable, then TMAO will acts as a small molecular crowding agent (26), that entropicallystabilizes the folded conformation(51). With these findings in mind, it is important to note that
our single-molecule thermodynamics results further support the established notion that, for 2°
structure formation, the osmolyte-induced change in ΔG° results from entropic terms (Figure6.6a). This notion suggest that the free energy change associated with expulsion of TMAO or

148

urea from the buried surfaces of 2° structure conformational transition is dominated by entropic
rewards, or costs, respectively.
To more quantitatively explore the osmolyte-dependence of the entropy change
associated with duplex formation and to reduce parameter correlation between ΔH° and ΔS°(52), the van’t Hoff plots are globally fit with common slopes and independent intercepts (Table
6.1). The resulting values of ΔS° are able to explain ≈ 80% of the osmolyte-dependence ofΔG°(295 K), which is entirely consistent with the notion of an entropic origin for urea-inhibitedor TMAO-facilitated duplex formation. An Eyring analysis of the temperature-dependent rate
constants (Supplemental Figure S6.1) unites the kinetic and thermodynamic origins of theseprocesses by demonstrating that most of the change in ΔS° is associated with the entropic
component of the unfolding free energy barrier (i.e., ΔS‡unfold), rather than the folding barrier(i.e., ΔS‡unfold). Stated more generally, TMAO stabilizes nucleic acid 2° structure formation by
decreasing kunfold, which occurs because of a decrease in the entropic reward associated with theunfolding barrier. Conversely, urea destabilizes 2° structure formation by increasing kunfold,which results from a more entropically rewarding unfolding barrier.
6.3.5

Osmolyte-Dependent Entropy Changes for the TL-TLR
When analyzed in the same fashion as the 8bp DNA duplex data, the temperature-dependent van’t Hoff analysis of the TL-TLR data makes it evident that addition of 1 M urea
primarily results in an entropic destabilization of this 3° interaction (Figure 6.6b). The resultsfrom the common-slope analysis reveal that the urea-induced changes to ΔS° account for ≈ 74%of the m-value at 295 K (Table 6.1). Furthermore, most of the change in ΔS° is associated withthe folding barrier (Supplemental Figure S6.2). This observation is entirely internally consistent
with the kinetic origin of urea-inhibited folding of the TL-TLR, which revealed that kfold is more149

prominently affected than kunfold (Figure 6.4c,d, triangles). Together, the kinetic andthermodynamic results suggest that urea destabilizes the TL-TLR by predominantly decreasing
kfold via an increase in the entropic cost associated with the folding barrier. That is to say, as the
RNA folds, urea is expelled from the buried nucleobase SASA, which is predominately an
entropically costly process.
Single-molecule urea-induced denaturation studies of an RNA pseudoknot(49) also
demonstrate a largely entropic destabilization, that results from a strong increase in kunfold rather
than a decrease in kfold, as observed for the TL-TLR. This observation suggests that individual
tertiary interactions likely have kinetics whose dependencies on urea are quantitatively different,
but that the destabilization by urea is always entropic in origin. Because of the canonical basepairing in both the RNA pseudoknot and the 8 bp DNA duplex, it is, perhaps, unsurprising thaturea predominately affects kunfold for these two folding transitions (RNA pseudoknot and DNA
duplex), as they are very similar with respect to changes in nucleobase SASA.
Although the kinetic origin of TMAO-facilitated formation of the TL-TLR isqualitatively similar to that of the 8bp DNA duplex (i.e., kfold increase and kunfold decreases), thethermodynamics of this process are noticeably different. Specifically, the presence of 1 M
TMAO significantly changes the ΔH° for the TL-TLR construct as evidenced by a more positive
slop on the van’t Hoff plot (Figure 6.6b). As a reminder, this effect is not observed in the 8bp
DNA duplex (Figure 6.6a). The fitted results reveal that both ΔH° and ΔS° decrease with theaddition of 1 M TMAO (Table 6.1), which mandates a thermodynamically distinct mechanism ofstabilization for the TL-TLR than that of the 8bp DNA duplex. One of the most pronounced
differences between the two folding transitions is the amount of phosphate SASA burial.
Therefore, it is reasonable to conclude that, for folding transition where there is a substantial

150change in phosphate SASA (e.g., 3° structure formation), the expulsion of TMAO from thesesurfaces is both exothermic and entropically costly. For the TL-TLR interaction, most of thechange in phosphate SASA occurs prior to formation of the transition state (Figure 6.4a,b andSupplemental Figure S6.2). Because the TMAO-induced decrease in ΔH‡fold is greater than thedecrease in TΔS‡fold (Supplemental Figure S6.2), addition of 1 M TMAO results in a netincrease in kfold at 295 K. The distinct difference in the thermodynamic origins for TMAOstabilized folding of the 8 bp DNA duplex and the TL-TLR construct highlights the two potential
modes by which TMAO can interact with nucleic acids: (i) strong unfavorable interactions
between TMAO and the phosphate backbone driven by both entropy and enthalpy that results instabilization nucleic acid folding transitions with substantial changes in phosphate SASA and (ii)weak unfavorable, predominately entropic interactions between TMAO and the nucleobase thatstabilize nucleic acid folding transitions with miniscule changes in phosphate SASA.
Although these single-molecule experiments explicitly demonstrate the kinetic andthermodynamic effects of TMAO and urea on two model constructs, it remains unknown how
general these observations are, both with respect to different osmolytes (e.g., glycine betaine,proline, sucrose, etc.) and different nucleic acid structural motifs (e.g., pseudoknots, Gquadruplexes, kissing loops, etc.). Further exploration of the role of osmolytes in nucleic acidfolding transitions, guided by single-molecule experiments that focus on the kinetics andthermodynamics origins, will help uncover a more comprehensive understanding of thecomplexities associated with biomolecules in cosolute-rich cellular environments.
6.4.

Conclusions
Single-molecule fluorescence techniques have been used to observe the effect ofstabilizing (TMAO) and destabilizing (urea) osmolytes on the folding/unfolding rate constants151for both an 8 bp DNA duplex and the GAAA tetraloop—tetraloop receptor motif. The results
demonstrate that, regardless of the type of the nucleic acid conformational transition (i.e.,
secondary or tertiary), TMAO always stabilizes nucleic acid structure formation by increasing
the folding rate constant and decreasing the unfolding rate constant. Conversely, urea always
destabilizes nucleic acid structure formation by decreasing the folding rate constant andincreasing the unfolding rate constant. The magnitudes of these osmolyte-induced changes to thefolding/unfolding rate constants are dependent on the amount and type (e.g., nucleobase) ofsolvent accessible surface area (SASA) that is buried for each of the conformational transition.
Complementary temperature-dependent single-molecule kinetic studies of the 8bp DNA
duplex reveal that osmolyte-induced stabilization or destabilization results almost entirely fromchanges to the entropic component of the free energy barrier for unfolding. For the TL-TLR
interaction, these studies demonstrate that urea destabilizes the structural equilibrium byincreasing the entropic penalty associated with the folding barrier. This is markedly different
than the thermodynamic origin of TMAO-facilitated formation of this structural motif, where an
increase in exothermicity overcomes an increase in entropic cost associated the folding freeenergy barrier. Together, these two effects results in a pronounced acceleration of the folding
rate constant.
As a whole, the experimental observations from this study represent a thoroughcharacterization of both the kinetic and thermodynamic origins of osmolyte-influenced nucleic
acid conformational transitions. Such a detailed characterization demonstrates how osmolytes
can be used as biochemical tools to map out changes in SASA during a particular folding
transition. Lastly, this work begins to unveil how small neutral organic molecules that are foundin cells and biological tissues may potentially influence the structural dynamics of functional

152

nucleic acids. Accordingly, future single-molecule efforts should be direct at studying the effectof other biologically relevant osmolytes like those that function in the renal medulla of thekidney(53) (e.g., glycine betaine, sorbitol, and taurine).
6.5

Experimental Methods

6.5.1

Nucleic Acids
The assembly of the smFRET DNA duplex construct has been described previously(33),

and needs only a brief explanation. This construct consists of three custom oligonucleotides(Figure 6.2a,b). Strand 1 (5#-biotin-(CA)8-Cy3-3#) has a biotin moiety for surface immobilization
and a donor fluorophore for fluorescence resonance energy transfer (FRET). This is annealed tostrand 2 (5#-TTG TTG GGT T(TG)8-3#) to form a 16 bp dsDNA scaffold with a 10 nt 5#
overhang. Strand 3 (5#-CCCAACCAA-Cy5-3#) contains a 3# acceptor dye, which, when present
in solution, can hybridize with the 5#overhang region (Figure 6.2b). Throughout such a
hybridization event, the Cy5 fluorophore of strand 3 remains in close proximity to the Cy3
fluorophore, allowing for efficient transfer of energy (EFRET ≈ 0.7) from the donor to theaccepter. After dissociation of strand 3 from the overhang region, diffusion rapidly increases the
distance between the donor and acceptor fluorophores (Figure 6.2a), which results in essentiallyzero energy transfer efficiency (EFRET ≈ 0). This gives rise to two possible FRET states for the
8bp DNA duplex construct—associated (Figure 6.2b, high EFRET) and dissociated (Figure 6.2a,low EFRET).
The GAAA tetraloop—tetraloop receptor (TL-TLR) model system is a modified version(51) of a previously described construct(39), where a synthetic polyethylene glycol (PEG) linker
is used to localize the GAAA tetraloop near its cognate receptor. This construct is built fromthree custom oligonucleotides (Figure 6.2c,d). The Cy3 labeled RNA that contains the GAAA

153

tetraloop (bold sequence) (5#-Cy3-GGC GAA AGC C-PEG6-CGU GUC GUC CUA AGU CGG
C-3#) is annealed to a Cy5 labeled RNA (5#-Cy5-GCC GAU AUG GAC GAC ACG CCC CUCAGA CGA GUG CG-3#) to form the 11 nt receptor domain (underlined sequence). This results
in a 14 nt 3# overhang that can hybridize to a complementary DNA containing a 5# biotin (5#Biotin-CGC ACT CGT CTG AG-3#) for surface immobilization. Once completely annealed, thisthree-piece construct serves as a FRET labeled, biotinylated TL-TLR construct. Formation ofthis interaction reduces the distance between the donor and acceptor fluorophores, which
increases the efficiency of energy transfer from EFRET ≈ 0.3 to EFRET ≈ 0.7. This results in two
distinct conformations for the TL-TLR construct—docked (Figure 6.2d, high EFRET) andundocked (Figure 6.2c, low EFRET).
6.5.2

Solution and Sample Preparation
Surface immobilization is accomplished via biotin-streptavidin chemistry. To attach thenucleic acids to a glass coverslip, the sample holder (10 µL) is consecutively flushed with 200µL of the following three solutions: (i) 10 mg/mL bovine serum albumin (BSA) containing 10%
biotinylated-BSA, (ii) 200 µg/mL streptavidin, and (iii) 100 pM biotinylated fluorescently
labeled nucleic acid. This procedure provides reproducible immobilization with surface densities
of approximately 1 molecule per µm2. Prior to imaging with the fluorescence microscope, thesample holder is flushed with 200 µL of an imaging buffer (50 mM hemisodium HEPES, 5.0
mM PCA, 10 mM KOH, 100 nM PCD, 2.0 mM Trolox, 0.1 mM EDTA at pH 7.5) containing thedesired concentrations of osmolytes (0-2 M) and KCl (either 50 mM or 100 mM). The enzymatic
oxygen scavenging system of PCD and PCA (54) is used to enhanced the photostability of thecyanine dyes. For experiments involving the 8 bp DNA duplex, the imaging buffer also contains

154

200 nM freely diffusing Cy5 oligonucleotide, which permits observation of the bimolecular
process.
6.5.3

smFRET Microscope
All smFRET experiments are performed on an inverted confocal fluorescence microscopewith a N.A. = 1.2 water objective (Figure 6.7).

Figure 6.7 Schematic representation of the inverted confocal fluorescence microscope used toacquire smFRET trajectories. Individual molecules are immobilized to the glass coverslip using
standard biotin-streptavidin chemistry. Excitation and emission is achieved using standard
epifluorescence techniques. Fluorescent photons are focused through a confocal pinhole before
being separated by color and polarization and directed onto 4 single photon avalanche
photodiodes. Upon detection, the arrival times of individual photons are recorded and sent to atime-correlated single-photon counting (TCSPC) router where they are used to generate
trajectories of donor and acceptor fluorescence.
A 20 MHz repetition rate, 532 nm laser beam overfills the limiting aperture of the optical system
resulting in a diffraction-limited laser spot at the focus of the objective, which is used to directly
excite the donor fluorophores of surface immobilized RNA constructs. Fluorescent photons are155

collected using a standard epifluorescence configuration before being separated by color (i.e.,
donor and acceptor) and polarization and directed onto four single photon avalanche photo
diodes (APDs). A Time-Correlated Single-Photon Counting (TCSPC) module records the arrivalof each detected photon. This information is used to construct time-traces of the donor (ID) andacceptor (IA) fluorescence intensity (Figure 6.3), which can readily be converted to FRET
trajectories using Eq. 6.1, where α is a term that corrects for various components of collection
efficiency (e.g., direct excitation, crosstalk see ref (35) for further information).
! ! !

!!"#$ = ! 1 − !! !! !!

! ! !

! !

1 + !! !!

! !

(Eq. 6.1)

Dwell times are determined from the FRET trajectories using a previously described
thresholding routine(34). Rate constants are extracted from cumulative distribution plots of dwell
times(41-43) by performing a linear least-squares fit to a single-exponential decay. Temperature
control of the sample is accomplished using commercially available stage and objective heaters,
as described previously(38).

156

6.6

References1.

Draper, D. E. A guide to ions and RNA structure, RNA, 2004, 10, 335-343.

2.

Klein, D. J.; P. B. Moore; T. A. Steitz. The contribution of metal ions to the structural
stability of the large ribosomal subunit, RNA, 2004, 10, 1366-1379.

3.

Kurz, M. Compatible solute influence on nucleic acids: many questions but few answers,
Saline Systems, 2008, 4, 6.

4.

Yancey, P. H.; M. E. Clark; S. C. Hand; R. D. Bowlus; G. N. Somero. Living with water
stress: evolution of osmolyte systems, Science, 1982, 217, 1214-1222.

5.

Record, M. T., Jr.; E. S. Courtenay; D. S. Cayley; H. J. Guttman. Responses of E. coli toosmotic stress: large changes in amounts of cytoplasmic solutes and water, Trends
Biochem. Sci., 1998, 23, 143-148.

6.

Yancey, P. H.; G. N. Somero. Counteraction of urea destabilization of protein structure
by methylamine osmoregulatory compounds of elasmobranch fishes, Biochem. J., 1979,
183, 317-323.

7.

Record, M. T., Jr.; E. S. Courtenay; S. Cayley; H. J. Guttman. Biophysical compensation
mechanisms buffering E. coli protein-nucleic acid interactions against changing
environments, Trends Biochem. Sci., 1998, 23, 190-194.

8.

Liu, Y.; D. W. Bolen. The peptide backbone plays a dominant role in protein stabilization
by naturally occurring osmolytes, Biochemistry, 1995, 34, 12884-12891.

9.

Timasheff, S. N. Control of protein stability and reactions by weakly interacting
cosolvents: the simplicity of the complicated, Adv. Protein Chem., 1998, 51, 355-432.

10.

Courtenay, E. S.; M. W. Capp; R. M. Saecker; M. T. Record, Jr. Thermodynamic
analysis of interactions between denaturants and protein surface exposed on unfolding:interpretation of urea and guanidinium chloride m-values and their correlation withchanges in accessible surface area (ASA) using preferential interaction coefficients andthe local-bulk domain model, Proteins, 2000, 4, 72-85.

11.

Courtenay, E. S.; M. W. Capp; C. F. Anderson; M. T. Record, Jr. Vapor pressure
osmometry studies of osmolyte-protein interactions: implications for the action of
osmoprotectants in vivo and for the interpretation of "osmotic stress" experiments invitro, Biochemistry, 2000, 39, 4455-4471.

12.

Lambert, D.; D. E. Draper. Effects of Osmolytes on RNA Secondary and Tertiary
Structure Stabilities and RNA-Mg2+ Interactions, J. Mol. Biol., 2007, 370, 993-1005.

13.

Lambert, D.; D. Leipply; D. E. Draper. The osmolyte TMAO stabilizes native RNA
tertiary structures in the absence of Mg2+: Evidence for a large barrier to folding from
phosphate dehydration, J. Mol. Biol., 2010, 404, 138-157.

157

14.

Lambert, D.; D. E. Draper. Denaturation of RNA secondary and tertiary structure byurea: simple unfolded state models and free energy parameters account for measured mvalues, Biochemistry, 2012, 51, 9014-9026.

15.

Spink, C. H.; N. Garbett; J. B. Chaires. Enthalpies of DNA melting in the presence ofosmolytes, Biophys. Chem., 2007, 126, 176-185.

16.

Hong, J.; M. W. Capp; C. F. Anderson; R. M. Saecker; D. J. Felitsky; M. W. Anderson;
M. T. Record, Jr. Preferential interactions of glycine betaine and of urea with DNA:
implications for DNA hydration and for effects of these solutes on DNA stability,
Biochemistry, 2004, 43, 14744-14758.

17.

Felitsky, D. J.; J. G. Cannon; M. W. Capp; J. Hong; A. W. Van Wynsberghe; C. F.
Anderson; M. T. Record, Jr. The exclusion of glycine betaine from anionic biopolymer
surface: why glycine betaine is an effective osmoprotectant but also a compatible solute,
Biochemistry, 2004, 43, 14732-14743.

18.

Bennion, B. J.; V. Daggett. The molecular basis for the chemical denaturation of proteins
by urea, Proc. Natl. Acad. Sc.i USA, 2003, 100, 5142-5147.

19.

Greene, R. F., Jr.; C. N. Pace. Urea and guanidine hydrochloride denaturation ofribonuclease, lysozyme, alpha-chymotrypsin, and beta-lactoglobulin, J. Biol. Chem.,
1974, 249, 5388-5393.

20.

Tanford, C. Isothermal Unfolding of Globular Proteins in Aqueous Urea Solutions, J.
Am. Chem. Soc., 1964, 86, 2050-&.

21.

Shelton, V. M.; T. R. Sosnick; T. Pan. Applicability of Urea in the Thermodynamic
Analysis of Secondary and Tertiary RNA Folding, Biochemistry, 1999, 38, 16831-16839.

22.

Canchi, D. R.; A. E. Garcia. Cosolvent Effects on Protein Stability, Annu. Rev. Phys.
Chem., 2013.

23.

Schwinefus, J. J.; M. J. Kuprian; J. W. Lamppa; W. E. Merker; K. N. Dorn; G. W. Muth.
Human telomerase RNA pseudoknot and hairpin thermal stability with glycine betaine
and urea: Preferential interactions with RNA secondary and tertiary structures,
Biochemistry, 2007, 46, 9068-9079.

24.

Cho, S. S.; G. Reddy; J. E. Straub; D. Thirumalai. Entropic Stabilization of Proteins by
TMAO, J. Phys. Chem. B, 2011, 115, 13401-13407.

25.

Wang, A. J.; D. W. Bolen. A naturally occurring protective system in urea-rich cells:
Mechanism of osmolyte protection of proteins against urea denaturation, Biochemistry,
1997, 36, 9101-9108.

26.

Pincus, D. L.; C. Hyeon; D. Thirumalai. Effects of trimethylamine N-oxide (TMAO) andcrowding agents on the stability of RNA hairpins, J. Am. Chem. Soc., 2008, 130, 73647372.

158

27.

Priyakumar, U. D.; C. Hyeon; D. Thirumalai; A. D. MacKerell. Urea Destabilizes RNA
by Forming Stacking Interactions and Multiple Hydrogen Bonds with Nucleic Acid
Bases, J. Am. Chem. Soc., 2009, 131, 17759-17761.

28.

Tinoco, I.; R. L. Gonzalez. Biological mechanisms, one molecule at a time, Gene Dev.,
2011, 25, 1205-1231.

29.

Roy, R.; S. Hohng; T. Ha. A practical guide to single-molecule FRET, Nat. Methods,
2008, 5, 507-516.

30.

Li, P. T.; J. Vieregg; I. Tinoco, Jr. How RNA Unfolds and Refolds, Annu. Rev. Biochem.,
2008, 77, 77-100.

31.

Bokinsky, G.; X. Zhuang. Single-Molecule RNA Folding, Acc. Chem. Res., 2005, 38,
566-573.

32.

Aleman, E. A.; R. Lamichhane; D. Rueda. Exploring RNA folding one molecule at atime, Curr. Opin. Chem. Biol. 2008, 12, 647-654.

33.

Dupuis, N. F.; E. D. Holmstrom; D. J. Nesbitt. Single-Molecule Kinetics Reveal CationPromoted DNA Duplex Formation Through Ordering of Single-Stranded Helices,
Biophys. J., 2013, 105, 756-766.

34.

Downey, C. D.; J. L. Fiore; C. D. Stoddard; J. H. Hodak; D. J. Nesbitt; A. Pardi. Metal
ion dependence, thermodynamics, and kinetics for intramolecular docking of a GAAA
tetraloop and receptor connected by a flexible linker, Biochemistry, 2006, 45, 3664–3673.

35.

Fiore, J. L.; J. H. Hodak; O. Piestert; C. D. Downey; D. J. Nesbitt. Monovalent andDivalent Promoted GAAA-Tetraloop–Receptor Tertiary Interactions from Freely
Diffusing Single-Molecule Studies, Biophys. J., 2008, 95, 3892-3905.

36.

Fiore, J. L.; E. D. Holmstrom; L. R. Feigland; J. H. Hodak; D. J. Nesbitt. The Role ofCounterion Valence and Size in the GAAA Tetraloop-Receptor Docking/Undocking
Kinetics, 2012, J. Mol. Biol., 2012, 423, 198-216.

37.

Fiore, J. L.; E. D. Holmstrom; D. J. Nesbitt. Entropic origin of Mg2+-facilitated RNA
folding, Proc. Natl. Acad. Sci. USA, 2012, 109, 2902-2907.

38.

Fiore, J. L.; B. Kraemer; F. Koberling; R. Erdmann; D. J. Nesbitt. Enthalpy-Driven RNA
Folding: Single-Molecule Thermodynamics of Tetraloop–Receptor Tertiary Interaction,
Biochemistry, 2009, 48, 2550–2558.

39.

Hodak, J. H.; C. D. Downey; J. L. Fiore; A. Pardi; D. J. Nesbitt. Docking kinetics andequilibrium of a GAAA tetraloop-receptor motif probed by single-molecule FRET, Proc.
Natl. Acad. Sci. USA, 2005, 102, 10505–10510.

40.

Holmstrom, E. D.; J. L. Fiore; D. J. Nesbitt. Thermodynamic Origins of Monovalent
Facilitated RNA Folding, Biochemistry, 2012, 51, 3732-3743.

159

41.

Blanco, M.; N. G. Walter. Analysis of complex single-molecule FRET time trajectories,
Methods Enzymol., 2010, 472, 153-178.

42.

Bartley, L. E.; X. W. Zhuang; R. Das; S. Chu; D. Herschlag. Exploration of theTransition State for Tertiary Structure Formation Between an RNA Helix and a Large
Structured RNA, J. Mol. Biol., 2003, 328, 1011–1026.

43.

Zhou, Y. J.; X. W. Zhuang. Robust reconstruction of the rate constant distribution using
the phase function method, Biophys. J., 2006, 91, 4045-4053.

44.

Sclavi, B.; S. Woodson; M. Sullivan; M. R. Chance; M. Brenowitz. Time-resolved
synchrotron X-ray "footprinting", a new approach to the study of nucleic acid structureand function: application to protein-DNA interactions and RNA folding, J. Mol. Biol.,
1997, 266, 144-159.

45.

Pan, J.; D. Thirumalai; S. A. Woodson. Folding of RNA involves parallel pathways, J.
Mol. Biol., 1997, 273, 7-13.

46.

Pace, C. N. The stability of globular proteins, CRC Crit. Rev. Biochem., 1975, 3, 1-43.

47.

Myers, J. K.; C. N. Pace; J. M. Scholtz. Denaturant m values and heat capacity changes:
relation to changes in accessible surface areas of protein unfolding, Protein Sci., 1995, 4,
2138-2148.

48.

Fiore, J. L.; E. D. Holmstrom; D. J. Nesbitt. Entropic Origin of Mg2+-Facilitated RNA
Folding, Proc. Natl. Acad. Sci. USA, 2012, 109, 2902-2907.

49.

Holmstrom, E. D.; D. J. Nesbitt. Single-molecule fluorescence resonance energy transfer
studies of the human telomerase RNA pseudoknot: temperature-/urea-dependent folding
kinetics and thermodynamics, J. Phys. Chem. B, 2014, 118, 3853-3863.

50.

Dalgarno, P. A.; J. Bordello; R. Morris; P. St-Pierre; A. Dube; I. D. Samuel; D. A.
Lafontaine; J. C. Penedo. Single-molecule chemical denaturation of riboswitches, Nucleic
Acids Res., 2013, 41, 4253-4265.

51.

Dupuis, N. F.; E. D. Holmstrom; D. J. Nesbitt. Molecular-crowding effects on singlemolecule RNA folding/unfolding thermodynamics and kinetics, Proc. Natl. Acad. Sci.
USA, 2013. (in Press)

52.

Cornish-Bowden, A. Enthalpy-entropy compensation: a phantom phenomenon, J. Biosci.,
2002, 27, 121-126.

53.

Burg, M. B. Coordinate regulation of organic osmolytes in renal cells, Kidney Int., 1996,
49, 1684-1685.

54.

Aitken, C. E.; R. A. Marshall; J. D. Puglisi. An Oxygen Scavenging System for
Improvement of Dye Stability in Single-Molecule Fluorescence Experiments, Biophys. J.,
2008, 94, 1826-1835.

160

Chapter 7

Single Molecule FRET Studies of the Human Telomerase RNA Pseudoknot:
Temperature/Urea Dependent Folding Kinetics and Thermodynamics*

7.1

Introduction
Conventional DNA replication presents a serious problem for the ends of linear

chromosomes. Each successive replication event results in the loss of genetic material at the endsof these long DNA duplexes, which has come to be known as the end replication problem (1).
One of nature’s solution to this problem is a ribonucleoprotein (RNP) complex, telomerase, thatfunctions to maintain the long-term fidelity of eukaryotic chromosomes(2,3). This task isaccomplished by processively adding multiple replicates of a species-specific DNA sequence tothe ends of the chromosomes, which results in the formation of telomeres(4). Not only does
repetitive elongation of telomeres buffer the natural shortening of chromosomes that occurs
during DNA replication, but the presence of the telomeres allows for recruitment of a number of
specific DNA binding proteins that are capable of protecting the chromosome ends from otherpotentially damaging cellular processes.
Proper telomere maintenance is essential for highly proliferative cells (e.g. stem cells)
and is partially responsible for cellar immortality. Unfortunately, the marvels of telomerase arealso beneficial for the highly proliferative constituents of cancerous tumors, with up-regulationof telomerase activity observed in > 85% of oncogenic cells(5). Indeed, the ability to repress
telomerase activity offers a broadly applicable target for cancer therapeutics, while the ability to!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!!
*
Adapted from: Holmstrom, Erik D; Nesbitt, David J. J. Phys. Chem. B. 2014, 118, 3853-3863!

161

enhance telomerase activity has applications in stem cell-based regenerative medicine. In orderfor medicinal applications to target telomerase activity more precisely, a deeper understanding of
the complete RNP is essential. Until recently, mechanistic information about telomerase
holoenzyme activity has proven quite challenging to obtain, though single-molecule studies ofmodel systems have recently started to elucidate some key structural and mechanistic aspects oftelomerase function (6-10). Specifically, these efforts have started to unveil details associatedwith the functional importance of the pseudoknot domain and the global conformation of the
RNA in catalytically active enzymes; however, much of the detailed biophysics of telomerase
activity still remains poorly understood.
What is known about telomerase is that the RNP consists primarily of two crucial
structural components, though additional species-specific components have been shown to berequired in various systems (11). The first is a protein subunit, telomerase reverse transcriptase(TERT), which is responsible for catalyzing the addition of telomeric repeats to the ends ofeukaryotic chromosomes(12). The second component is telomerase RNA (TR), which, among
other things, provides the template for this RNA-dependent DNA synthesis (12). One of the few
highly conserved domains within TR is the core domain, which is essential for both RNP
assembly and activity (13-19). It has been shown that within this domain there is a functionallycritical region that forms a pseudoknot motif. In general, a pseudoknot is formed when the
nucleotides in the loop of an RNA hairpin make stable, typically canonical, base pairing
interactions with the free nucleotides of a single stranded region elsewhere in the RNA. The
pseudoknot base pairing interactions (P3) found in human TR (hTR) are formed, in part, between
the nucleotides in the loop of the P2 hairpin and the single stranded nucleotides of J2a/3 (Figure7.1a). A number of NMR experiments have determined the molecular structure of a minimal!

162

wild-type (WT) hTR pseudoknot (Figure 7.1b) and various mutants(20-24). These structuralexperiments, together with recent single-molecule experiments(25,26) and computational folding
models (27,28), indicate that the WT pseudoknot exists in equilibrium between a partiallyunfolded hairpin and a folded pseudoknot (PK) with a stable and conserved triple helix (Figure7.1c).

Figure 7.1 Details of the human telomerase RNA pseudoknot a) Secondary structure diagramsof the minimal pseudoknot motif in a partially unfolded (hairpin) conformation and a folded(pseudoknot) conformation. The GC!AG mutation at the orange nucleotides in the psuedoknot
region corresponds to the genetic disorder dyskeratosis congenita. Green (Cy3) and red (Cy5)stars indicate the location of two fluorophores, where size of the stars reflect FRET efficiency
between the two dyes and the black vertical bar represents the ligation site. b) Model solution
structure of the minimal pseudoknot motif, with the same color scheme as the secondary
structure diagram. c) Schematic equilibrium between the pseudoknot and hairpin conformations.
Such detailed investigations of the pseudoknot region of hTR have been used in conjunction withexisting mutational studies to develop several models for the functional importance of thisstructural motif. To date, however, substantial accumulation of evidence in support of (oragainst) any of these model predictions has remained elusive, which therefore represents
substantial motivation for the present work.

!

163

In this study, single-molecule fluorescence resonance energy transfer (smFRET)
techniques(29,30) are used to further characterize the kinetics and thermodynamics of two
minimal hTR PK constructs: (i) the wild-type (WT hTR PK) and (ii) a mutant thereof that is
associated with the genetic disorder dyskeratosis congenita (DKC hTR PK). These smFRET
experiments provide a remarkably quantitative methodology for observing molecular dynamics
of biomolecules. Single-molecule techniques are extremely advantageous for kinetic studies ofRNA folding in that they are able to resolve conformational dynamics unobservable in more
traditional ensemble measurements. The ability to measure the kinetics of RNA folding at
equilibrium and in real-time significantly increases the amount of information attainable from theexperimental system(31-33). Furthermore, the temperature-dependence of these smFRET studies
can be exploited to extract the enthalpic and entropic components associated with freeenergies(34-37). Lastly, experiments that utilize urea as a chemical denaturant provide important
quantitative information about the free energy stability of the pseudoknot motif (36,38). The
results of this study and the associated implications are discussed with regard to: (i) the folding
pathway of the minimal RNA pseudoknot motif, (ii) the use of urea as a general tool for probing
the single molecule kinetics and thermodynamics of RNA folding, and (iii) the molecularfunction and various disease states of telomerase.
7.2

Results

7.2.1

Freely Diffusing Burst Measurements
In the standard fluorescence-imaging buffer (see materials and methods), the wild-type(WT) human telomerase RNA pseudoknot (hTR PK) appears to exist in a single high FRET
conformation (EFRET = 0.95(1), FWHM = 0.11(1))(Figure 7.2a, black). This EFRET corresponds
to a length of R ≈ 32(1) Å which, when one considers the flexible carbon linkers (≈ 5 Å)!

164

connecting each of the two fluorescent dyes to the RNA, is consistent with the modeled distance(≈ 22 Å, ref (20)) between the two nucleotides nearest to the fluorophores. Therefore, theobserved high FRET state is the result of a stably folded pseudoknot.

Figure 7.2 Freely diffusing FRET histograms of a) the wild-type human telomerase RNA
pseudoknot and b) the dyskeratosis congenita mutant of the human telomerase RNA pseudoknot.
The urea titrations demonstrate strong shifts in the pseudoknot-hairpin equilibrium behavior forboth constructs.
This is in stark contrast to the equilibrium distributions of FRET states associated with
the DKC hTR PK (Figure 7.2b, black). Under the same experimental conditions, the dyskeratosis
congenita (DKC) hTR PK adopts two conformations: (i) a high FRET conformation (EFRET =0.94(2), FWHM = 0.14(2)), indistinguishable from the pseudoknot structure for WT hTR PK and(ii) a low FRET conformation (EFRET = 0.07(1), FWHM = 0.19(1)), indicative of a substantial
increase in donor-acceptor distance. From the 1/R6 dependence of FRET efficiency on distance,
this low FRET value corresponds to ≈ 80(2) Å of separation between the two dyes and is indeed
consistent with the hairpin conformation.

!

165

To better assess the stability of the WT hTR PK and to explore other partially folded or
unfolded conformations, freely diffusing experiments have been performed as a function of urea
concentration (Figure 7.2a). Upon addition of 7.7 M urea, a second conformation with low FRET
(EFRET = 0.11(1), FWHM = 0.20(1)) begins to appear, which is essentially indistinguishable fromthe low FRET conformation of the DKC hTR PK under non-denaturing conditions (Figure 7.2a,blue and Figure 7.2b, black). This suggests that the low FRET state for WT hTR PK in thepresence of high urea concentrations is almost identical to the extended hairpin structure that the
DKC hTR PK occupies under non-denaturing conditions. For the WT hTR PK, the relative
population of molecules in the hairpin conformation continues to increases with increasing
denaturant concentration up to the solubility limit of urea (≈ 10.3 M). This same trend isapparent for the DKC hTR PK, however it occurs over a much more dilute range of urea
concentrations (Figure 7.2b). Both smFRET constructs are able to reversibly and completely
redistribute between the low and high FRET states depending upon urea and monovalent cation
concentration. Such behavior is consistent with a simple, two-state, unimolecular equilibrium
between the pseudoknot and hairpin conformations.
An alternatively labeled RNA construct (WTalt hTR PK) has been designed to further
validate that the WT hTR PK exists in a hairpin-pseudoknot equilibrium under all urea
concentrations (Supplemental Figure S7.1a). For this construct, the fluorophore labelingspecifically probes disruption and formation of the P2 region of the pseudoknot, rather than the
P3 region sampled with the conventional labeling scheme. The WTalt hTR PK exists solely in ahigh FRET conformation at both 0 M urea (EFRET = 0.95(1), FWHM = 0.09(1)) and 10.3 M urea
(EFRET = 0.95(1), FWHM = 0.13(1)), which confirms that the observed dynamics in the!

166conventional construct are indeed the result of disruption and formation of the P3 region of theWT hTR PK (Supplemental Figure S7.1b).

Supplemental Figure S7.1 a) Details of the alternative labeling scheme for the wild-type humantelomerase RNA pseudoknot secondary structure. The minimal pseudoknot motif is in a partiallyunfolded (hairpin) conformation and a folded (pseudoknot) conformation. The green (Cy3) andred (Cy5) stars indicate the location of the two fluorophores, while the size of the stars represents
the FRET efficiency between the two dyes. The grey vertical bar represents the location of theligation site (see Section 7.5.1 for details) b) Freely diffusing FRET histograms of alternate wildtype human telomerase RNA pseudoknot. The nearly identical FRET values at high and low urea
concentrations confirm that: (i) the P2 region of the pseudoknot remains intact under all
experimental conditions and (ii) the dynamics associated with the conventional constructs resultsfrom formation/disruption of the P3 region of the pseudoknot.
Given that a unimolecular equilibrium can be used to accurately describe the hairpinpseudoknot transition of both the WT and DKC hTR PK constructs used in this study, the ratioof the relative population of the pseudoknot and hairpin conformations yields the equilibrium
constant (Keq = [pseudoknot]/[hairpin]) for the two species (Figure 7.1c). The equilibrium
constant can easily be converted to the standard state Gibb’s free energy change associated withtransitioning between the two conformations (i.e., ΔG° = −RTln[Keq]). As previously observed in!

167other RNA systems(36,38,39), the free energy change depends linearly on the urea concentration(Figure 7.3). Although the slopes (m-values) for the two constructs are not identical (seeDiscussion), the pronounced linearity of both plots at high and low urea concentrations makes it
reasonable to assume linear behavior across all urea concentrations. As a result, a simple linear
extrapolation to 0 M urea should yield the ΔG°(295 K) values associated with hairpin-topseudoknot transitions under non-denaturing conditions (ΔG°DKC = 0.26(4) kcal/mol, ΔG°WT =−4.2(2) kcal/mol).

Figure 7.3 Urea dependence of the free energy change for forming the pseudoknot from thehairpin conformation for the wild-type human telomerase RNA pseudoknot (grey) and thedyskeratosis congenita mutant (orange) of the human telomerase RNA pseudoknot. Linear
extrapolation to 0 M urea yields the 4.5(3) kcal/mol differential folding free energy between the
WT and mutant pseudoknot under non-denaturing conditions.
7.2.2

Surface Immobilized Images
ALEX-filtered, raster-scanned surface images of immobilized molecules qualitatively

recapitulate the results of the freely diffusing experiments (Figure 7.4). Under non-denaturingconditions, the WT hTR PK molecules exist entirely in the high FRET (pseudoknot)
conformation (Figure 7.4a), while the DKC hTR PK molecules are distributed between the two
conformations, with the low FRET (hairpin) state being slightly more populated (Figure 7.4c).

!

168

Upon closer inspection, however, the surface immobilized images provide partial information on
the lifetime associated with the pseudoknot and hairpin conformations. Specifically, theuniformity of color for each diffraction-limited spot (Figure 7.4a,c,d) suggests that dwell times
for the pseudoknot and hairpin conformations are, on average, longer than the ≈ 10 s time scale
sampled in rastering across each of the ≈ 10 horizontal rows of pixels that comprise a single
fluorescent spot.

Figure 7.4 Images of surface immobilized molecules for the wild-type a,b) and dyskeratosis
congenita c,d) pseudoknots. Each diffraction limited fluorescence spot represents the location of
an individual dually labeled RNA molecule. The urea dependence of the images yields kinetic
information about the two different pseudoknot constructs (see Section 7.2.2 for details).
This contrasts subtly but quite significantly with images for WT hTR PK at 10.3 M urea (Figure7.4b), where individual fluorescent spots exhibit both red and green contributions, but only withcomplete rows of either fully red or green pixels. Such color continuity in the horizontal but notnecessarily vertical dimension suggests that switching must take place on a time-scale that is i)clearly faster than 10 second overall scan time, but apparently also ii) slower than the ≈ 10 ms it
takes to scan across ≈ 10 horizontal pixels within a single row for a given fluorescence spot.

!

1697.2.3

Surface Immobilized Time Trajectories
To obtain more quantitative kinetic information about the rate constants for transitioning

between the pseudoknot and hairpin conformations, donor-acceptor fluorescence time
trajectories are acquired for individual surface immobilized molecules (Figure 7.5a,c).

Figure 7.5 Surface immobilized folding and unfolding kinetics for the a) wild-type pseudoknot
and c) the mutant human telomerase RNA pseudoknots under denaturing and non-denaturingconditions, respectively. Rate constants are determined from exponential fits to cumulative
distribution plots of dwell times for the b) wild-type and d) dyskeratosis congenita pseudoknots.

!

170

The fluorescence trajectories can easily be converted to FRET trajectories that cleanly
demonstrate the two-state behavior for both RNAs, with FRET values consistent with the freely
diffusing experiments above. For such a two-state equilibrium system, the dwell times associatedthe pseudoknot and hairpin states should be distributed exponentially, with characteristic time
constants corresponding to rate constants for unfolding (kunfold, pseudoknot → hairpin) or folding(kfold, pseudoknot ← hairpin) (Figure 7.1c). This expectation is nicely confirmed for both WT(Figure 7.5b) and DKC (Figure 7.5d) RNAs under denaturing and non-denaturing conditions,
respectively. Clearly, both pseudoknot and hairpin dwell times are distributed exponentially withwell-defined single exponential rate constants.
The FRET time trajectories for the WT hTR PK under non-denaturing conditions are
overwhelmingly dominated by the high FRET state. Given the finite observation window offluorescent molecules prior to photobleaching (≈ 100 sec), only a few conformational transitions
have been observed under non-denaturing conditions. This prevents quantitative determination of
dwell times and therefore rate constants associated with the WT construct under non-denaturingconditions. However, the fact that very few time trajectories exhibit even a single unfolding
event during a 100 s photobleaching lifetime suggests an upper-limit for the unfolding rate
constant of ≈ 0.01 s-1.
To achieve a better understanding of the WT hTR PK folding kinetics, surface
immobilized time trajectories have been acquired under denaturing conditions (Figure 7.5a).
Interestingly, high urea concentrations strongly increase the unfolding rates without substantiallyinfluencing the folding rates (Table 7.1a, Figure 7.6a). From the kinetics experiments, it is also
possible to readily extract equilibrium constants for the pseudoknot-hairpin system (Keq =kfold/kunfold), based on the assumption of a two-state system. As expected, the surface immobilized!

171

equilibrium constant data, derived from folding/unfolding kinetics, are in excellent agreementwith the freely diffusing experiments (Figure 7.3). Such agreement between surface immobilized
and freely diffusing data sets provides additional evidence to support that surface immobilization
does not perturb the conformation dynamics for either the WT or DKC constructs.

Figure 7.6 Log-linear plot of the urea-dependent folding and unfolding kinetics for the a) wildtype and b) dyskeratosis congenita pseudoknots. For both constructs, the unfolding rate constantis substantially more sensitive to addition of urea than the folding rate constant, which suggeststhat most of the change in base solvent accessible surface area (SASA) occurs after formation of
the transition state. Lines indicate trends and are not actual fits to the data.
The linear dependence of the folding free energy change on urea concentration (Figure7.3) can be used to develop a more quantitative understanding for the folding kinetics of the WT
hTR PK under non-denaturing conditions. Specifically, the extrapolation of ΔG° to 0 M urea in
Figure 7.3 yields ΔG°WT(295 K) = -4.2(2) kcal/mol, which would imply Keq = kfold/kunfold ≈ 1100
and thus a highly stable WT pseudoknot. Experimentally, the folding rate constant is observed tobe mostly insensitive urea concentration, which would imply any urea-dependence of ΔG° is due
primarily to changes in the unfolding rate constant. Therefore, a urea-independent folding rate
constant of ≈10 s-1 from Figure 7.6a would suggest an unfolding rate constant of ≈ 0.009 s-1 at 0!

172

M Urea (Table 7.1a), which is certainly consistent with the ≈ 0.01 s-1 upper limit established byphotostability of the present donor-acceptor FRET pair.

Table 7.1 Rate and equilibrium constants for the a) wild-type and b) mutant forms of the humantelomerase RNA pseudoknot under varying urea concentrations. For both RNA pseudoknot
constructs, the rate constant for forming the pseudoknot from the hairpin conformation is largely
insensitive to urea concentration.
In contrast with the WT construct, each trajectory for the DKC hTR PK construct under
non-denaturing conditions undergoes multiple conformational transitions between the two FRET
states (Figure 7.5c), which permits accurate determination of dwell times and rate constants(Figure 7.5d). As was the case for the WT hTR PK, addition of urea has only a small effect on
kfold while substantially increasing kunfold (Table 7.1, Figure 7.6b). It is worth noting that theabove kinetic measurements for the DKC hTR PK under non-denaturing conditions serve as avaluable benchmark for the corresponding rate constants for WT hTR PK extrapolated to 0 M
urea. Interestingly, such a comparison reveals that the net 4.5(3) kcal/mol destabilization of the!

173mutant DKC pseudoknot is achieved by a combination of much slower folding (≈ 400x) and only
slightly faster (≈ 5x) unfolding rate constants (Table 7.1).
7.2.4

van’t Hoff Thermodynamics
Via direct measurement of the equilibrium constant, these single molecule studies havethus far focused on standard Gibbs free energy change (ΔG°) as the primary, experimentally
determined, thermodynamic parameter. However, temperature-dependent information about theequilibrium constant can be used to deconstruct Gibbs free energy changes into enthalpic (ΔH°)
and entropic (ΔS°) components. Furthermore, the observation that ΔG° is linearly dependent onurea concentration makes it reasonable to assume that the two additive components of ΔG° =
ΔH° -TΔS° might also be linearly dependent on urea, which facilitates additional thermodynamic
comparisons between the wild type and mutant RNA constructs.
In freely diffusing experiments, the equilibrium constant for the WT hTR PK has beenmeasured at temperatures ranging from 20.0(1) °C to 33.0(1) °C for three different urea
concentrations (7.7 M, 8.4 M, and 9.1 M). At all urea concentrations, increasing the temperature
results in unfolding of the pseudoknot, thus reducing the equilibrium constant. As shown in
Figure 7.7a, a van’t Hoff plot (ln[Keq] vs 1/T) of the data is used to infer the entropy (ΔS° =intercept × R) and enthalpy (−ΔH° = slope × R) change associated with pseudoknot formation. It
is interesting to note the nearly identical slopes for each of the plots, i.e., enthalpy changes (ΔH°)
associated with pseudoknot formation are largely insensitive to urea. Along with morequantitative results from a non-linear least squares fit (Table 7.2a), these data sets provide clear
evidence that the addition of urea primarily decreases the folding entropy (ΔΔS° < 0), with anegligible effect on the folding enthalpy (ΔΔH° ≈ 0). To further break parameter correlation andthereby more precisely examine the dependence of ΔS° on urea, these van’t Hoff data sets have!

174

been re-fitted with a common enthalpic slope. As shown in Figure 7.7b, the resulting entropychanges are linearly dependent on urea concentrations and predict ΔS° = -250.2(6) cal/molK at 0
M urea. In conjunction with the urea-independent ΔH° = -78.2(6) kcal/mol, this limiting ΔS°
value corresponds to ΔG°(295 K) = -4.4(6) kcal/mol under non-denaturing conditions.

Figure 7.7 Single-molecule van’t Hoff analyses. a) A plot of the temperature-dependence of theequilibrium constant of the wild-type human telomerase RNA pseudoknot at three urea
concentrations. The identical slopes indicate a urea-independent folding enthalpy, while theconstant vertical offset indicates a urea-dependent entropy. b) A plot of the urea-dependent
entropy suggests a linear dependence of folding entropy on urea concentration. c) van’t Hoffanalysis of the mutant pseudoknot under non-denaturing conditions. All entropy and enthalpy
changes can be found in Table 7.2.

!

175

It is worth noting that this is experimentally indistinguishable from the ΔG°(295 K) = -4.2(2)kcal/mol value obtained from extrapolation of the freely diffusing data (Figure 7.3a), which
provides additional support for analyses of this conformational transition based on a linear
dependence of free energy and entropy on urea.

Table 7.2 Thermodynamics parameters from the single-molecule van’t Hoff analyses. a)
Entropy and enthalpy changes for the wild-type human telomerase pseudoknot at various urea
concentrations. ΔH° and ΔS° are (i) insensitive or (ii) linearly dependent on urea concentration,
respectively. Together they predict folding thermodynamics under non-denaturing conditions. b)
Enthalpy and entropy changes for the dyskeratosis congenita mutant form of the humantelomerase RNA pseudoknot under non-denaturing conditions.
To enable thermodynamic comparison between the two RNA pseudoknot constructs
under non-denaturing conditions, temperature-dependent equilibrium constants for DKC hTR PK
have also been measured. The resulting van’t Hoff plot (Figure 7.7c) predicts ΔS° = -129(5)cal/mol/K and ΔH° = -37(2) kcal/mol at 0 M Urea (Table 7.2b). Such a van’t Hoff analysis of the
DKC hTR PK data predicts ΔG°(295 K) = 1(2) kcal/mol at 0 M Urea, which is also consistent
with the ΔG°(295 K) = 0.26(5) kcal/mol value obtained from the freely diffusing data (Figure!

176

7.3b). Lastly, comparison of the WT and DKC constructs reveals that pseudoknot formation in
the mutant is both half (≈ 47%) as exothermic and half (≈ 51%) as entropically costly, whichmakes formation of the WT pseudoknot more favorable thermodynamically.
7.3

Discussion
The above studies demonstrate the well-behaved, two-state nature of both the wild-type(WT) and dyskeratosis congenita (DKC) variants of the minimal hTR PK, which is consistent
with recent smFRET findings (26). As a function of experimental conditions, both constructs areable to switch completely and reversibly between readily distinguishable low and high FRET
states. This observation supports the existence of a two-state pseudoknot-hairpin unimolecularequilibrium for both the WT and DKC constructs (23). Implications of the kinetic andthermodynamic parameters associated with this equilibrium are discussed in detail below.
7.3.1

Folding Free Energies
For solution conditions used throughout these studies, which appropriately mimic

physiological concentrations of free monovalent ion, the two RNA constructs exhibit vastlydifferent ΔG°(295 K) values for pseudoknot formation. The free energy of folding for the WT
hTR PK is -4.2(2) kcal/mol, as determined by a linear extrapolation to 0 M urea (Figure 7.3a),which should be compared to 0.26(5) kcal/mol for the DKC variant. Such a substantial stability
difference between the WT and DKC constructs (ΔΔG°(295K) = 4.5(3) kcal/mol) is qualitatively
consistent with existing thermal denaturation studies (6.6 kcal/mol) and theoretical calculations(6 kcal/mol) (23), especially accounting for differences in solution conditions (e.g. salt, pH,temperature) associated with each value. Furthermore, the overall sign of this shift is in the
expected direction, with the 2 nt non-complementary substitution resulting in less favorable
formation of the pseudoknot.

!

177

It is instructive to explore how ΔΔG°(295 K) between the WT and DKC constructs(4.5(3) kcal/mol) manifests itself in the kinetics of the hairpin-pseudoknot unimolecularequilibrium. As noted previously, the presence of the DKC mutation results in a large reduction(≈400-fold) and yet only a modest increase (≈5-fold) in the folding and unfolding rate
constants, respectively. In conjunction with the overall ΔΔG°, such kinetic information can beused to build a simple free energy reaction coordinate for the hairpin-pseudoknot folding
transition (Figure 7.8).

Figure 7.8 Free energy landscape for formation of the pseudoknot from the hairpin
conformation. Secondary structure diagrams for the wild-type human telomerase RNA
pseudoknot are shown, including the predicted structure of the transition state where only a few
nucleotides from the P3 region of the pseudoknot have formed (see Section 7.3.1 for details).
Specifically, the measured rate constants data are used to estimate the free energy barrier forfolding from transition state theory (Eq. 7.1), which would be consistent with a 15-20 kcal/mol
transition state barrier along the reaction coordinate for typical attempt frequency of ν ≈ 1013 s-1.
k

ΔG‡ = −RT ln ν!

178(Eq. 7.1)

It is worth noting, however, that ΔG‡ depends only logarithmically on ν; thus, any differentialchanges in free energy barriers predicted from Eq. 7.2
ΔΔG‡ = RT ln

kWT
kDKC

(Eq. 7.2)

should specifically remain completely independent of any such ad hoc and/or estimated choices
for the prefactor, ν. From this expression, the changes associated with the folding and unfolding
rate constants for the DKC mutant correspond to a 3.6 kcal/mol increase and a 0.9 kcal/mol
decrease in the folding and unfolding free energy barriers, respectively.
Furthermore, we can extract additional physical insights into these changes in barrierheight from a Φ-analysis (36,40). Specifically, for a given pair of modifications (e.g., DKC and
WT), Φ represents the fraction of the free energy barrier change for folding (ΔΔG‡fold) that
results per unit change in the overall equilibrium stability (ΔΔG°) as shown in Eq. 7.3.
∆∆G ‡

295K

Φ= ∆∆!fold! 295K(Eq. 7.3)

A Φ-analysis for the DCK and WT hTR pseudoknots demonstrates that 80% of the ΔΔG°(295 K)
comes from decreasing the DKC folding rate constant. This implies that the WT nucleotides atthe positions associated with the DKC mutation (i.e., G108 and C109) play an important role inlowering the free energy transition state barrier for folding. Furthermore, the results of the Φanalysis can be used to provide insights into the structure of the transition state for thisconformational transition. Specifically, this strongly suggests that the WT hTR PK constructs atthe transition state already have well-formed stabilizing contacts (e.g., base stacking andhydrogen bonding) at nucleotides G108 and C109.

!

179

7.3.2

Urea-dependent Free Energies and Rate Constants
Denaturation of nucleic acids is thought to occur because of favorable interactions

between urea and the base “solvent accessible surface area” (SASA)(41). For RNA in a
conformational equilibrium, the presence of urea preferentially stabilizes the structure with themost base SASA—typically unfolded and/or less compact. This manifests itself as a linear
dependence of the folding free energy change on urea concentration (∂ΔG°/∂[urea]), which isoften referred to as the m-value (38,42). Consequently, one would expect the m-value for RNA
folding to be positively correlated with the amount of base SASA buried due to a folding
transition, as has been shown in several studies(38,39). It is therefore interesting to note that them-values for the WT and DKC constructs differ in a counterintuitive direction. Specifically, due
to two fewer base pairing interactions in the P3 region, pseudoknot formation might be
anticipated to lead to less base SASA burial in DKC vs. WT hTR PK. As a result, one would
expect WT hTR PK folding to be more sensitive to urea, when in fact it is nearly half as sensitive(WT: m = 0.44(2) kcal/mol/M, DKC: m = 0.81(6) kcal/mol/M). The precise origin of thisdifference is not well understood. However, these results certainly indicate unexpecteddifferences in base SASA burial, which could potentially result from structural differences
between: (i) the WT and DKC hairpin conformations and (ii) the WT and DKC pseudoknot
conformations that may be related to the formation of base triples in the WT hTR PK. Together
these structural differences could give rise to larger base SASA burial in the DKC hTR PK,
which would be consistent with the experimentally observed m-values.
This m-value analysis can be similarly applied to the folding kinetics, by assuming thatthe urea-dependence of the rate constants is also related to the amount of base SASA burial
associated with formation of the transition state that separates the two conformations(36). For!

180both the hTR PK constructs, the very modest urea-dependence of the folding rate constantsuggests that there is little base SASA burial associated with forming the transition state from thehairpin conformation. In conjunction with the observation that nucleotides 108 and 109 are
important for fast folding, the urea-independent folding rate constant for the hTR PK imply atransition state where only the last two base pairs in P3 (e.g., G108 and C109) are formed,
leaving the remainder of the bases in J2a/3 solvent exposed (Figure 7.8). Minimal burial of base
SASA resulting from formation of the transition state from the hairpin conformation provides therational for a largely urea-insensitive folding rate constant, as observed experimentally.
Accordingly, most of the total base SASA burial takes place after the transition state duringformation of the remainder of the P3 pseudoknot, which gives rise to a substantial dependence of
the unfolding rate constant on urea concentration. Interestingly, similar conclusions have been
made in quite different RNA constructs (36,43), which: (i) suggest that urea may generally
destabilize all RNA tertiary folding transitions by selectively accelerating the unfolding rate
constant and (ii) support the notion that many transition states for RNA tertiary folding may be
conformationally aligned, yet still devoid of significant base SASA burial (e.g., minimalhydrogen-bonding and base stacking).
7.3.3

Urea-Dependent Entropies and Enthalpies
The measured urea-independent enthalpy for WT pseoduknot formation (ΔH° = -78.2(6)kcal/mol) is remarkably close to both theoretical calculations (-78.9 kcal/mol (44)) and previous
experimental observations using ensemble techniques (-72.7 kcal/mol (22,23)). Such quantitative
agreement serves as strong support for the use of urea as a valuable tool to study highly stable
RNAs at the single-molecule level under denaturing conditions. Additionally, van’t Hoff analysis
as a function of urea yields new insights into the thermodynamic origin of urea-induced!

181denaturation of the hTR PK. For example, the near complete insensitivity of ΔH° to urea
provides experimental evidence that destabilization associated with urea is predominantly due tochanges in ΔS°. This entropic origin of urea destabilization, together with the observation thaturea has little effect on the folding rate constant, suggests that the majority of the denaturing
power of urea results from increasing the entropic reward associated with forming the transition
state from the pseudoknot conformation. Simply stated, urea destabilizes the WT hTR PK bymaking unfolding more entropically beneficial. This is a surprising observation, given that the
proposed mechanism for urea denaturation of nucleic acids results from favorable hydrogenbonding between urea and the base SASA of RNA. The lack of any substantial enthalpic
component to urea denaturation therefore must imply nearly perfect cancellation of any
differential exothermicity between the competing urea-base and base-base interactions.
Additionally, the determination of a urea-independent enthalpy change and a linear ureadependent entropy change make it feasible to compare the folding thermodynamics between the
WT and DKC hTR PK constructs used throughout this work (Figures 7.7a,c, and Table 7.2).
One additional surprising observation is that the increase in folding free energy (ΔΔG° =4.5(3) kcal/mol) associated with the WT vs. DKC mutant results from a fortuitous nearcancellation between much larger changes of both the folding enthalpy (ΔΔH° ≈ 41 kcal/mol)and entropy (ΔΔS° ≈ 121 kcal/mol/K, TΔΔS° ≈ 37 kcal/mol). An approximately 50% reductionin the enthalpic and entropic components of pseudoknot formation is certainly unexpected, but
clearly indicates that substantial thermodynamics differences exist between the WT and the DKC
hTR PK. These large entropic and enthalpic differences may be related to the vastly different mvalues associated with the two constructs. As mentioned previously, it is likely that the WT and
DKC constructs have different pseudoknot conformations. Specifically, the DKC hTR PK may!

182

not be able to from the well-defined triple-helical structure of the WT sequence. If this were thecase, then pseudoknot formation in the DKC construct would lack the additional exothermicity
as well as the entropic cost associated with forming the five base triples within the WT triple
helix, resulting in the large ΔΔH° and ΔΔS° values that are observed experimentally.
Unfortunately, differentiation between these two types of folding conformations (i.e., pseudoknot
vs. triple-helix) is not possible with the current FRET constructs, but clearly identifies aninteresting direction for future structural efforts.
7.4

Conclusions
Single molecule FRET experiments have been used to cleanly observe folding transitions

in a minimal RNA pseudoknot (PK) designed to mimic the wild-type motif within the humantelomerase RNA. These experiments are compared and contrasted with experiments conducted
on a second minimal human telomerase RNA pseudoknot that contains a 2 nt noncomplementary substitution mutation implicated in the telomerase-associated genetic disorder—
dyskeratosis congenital (DKC). These experiments have demonstrated that the wild-type
pseudoknot is substantially more stable (ΔΔG = 4.5(3) kcal/mol) than the mutant. Singlemolecule kinetic studies of these constructs have identified the kinetic origin of this differential
stability as being the result of: (i) a substantially reduced folding rate constant and (ii) a
moderately increased unfolding rate constant, both of which contribute to destabilization of the
fully folded mutant pseudoknot conformation. With the help of urea-induced denaturation,
smFRET experiments have been used to elucidate the free energy landscape and various
structural aspects of the folding reaction coordinate. Lastly, single-molecule van’t Hoff analyses
highlight a potential folding pathway wherein the mutant pseudoknot is unable to form thefunctionally important triple-helix structure known to exist in the wild-type RNA. Together,!

183

these results suggest that the disease mechanism associated with this particular genetic mutationis related to a combination of: (i) differential stabilities of the wild-type and mutant pseudoknot
conformations, (ii) differential folding/unfolding kinetics associated with the pseudoknot-hairpin
unimolecular equilibrium for the wild-type and mutant RNAs, and (iii) the potential lack of the a
RNA triple helix in the mutant pseudoknot. Precise determination of the detailed mechanism of
telomerase and the associated disease states would benefit substantially from extensions of thesesingle-molecule experiments on minimal hTR pseudoknots to more holoenzyme-like systems
that more closely recapitulate telomerase activity and function.
7.5

Experimental Methods

7.5.1

Construct Design
To generate suitable FRET labeled wild-type (WT hTR PK) and mutant (DKC hTR PK)

psuedoknot constructs for single molecule studies, we exploit the powerful and increasingly
common technique of RNA ligation (45-48). For the wild type construct, two syntheticallymodified oligonucleotides—strands 1 and strand 2—are purchased commercially (Integrated
DNA Technologies, Inc). Strand 1 contains both a 5) biotin moiety for surface immobilization as
well as a internal amino modified dT (dT*) at position 106 (numbering from full lengthhTR(13)) for fluorescent labeling: (5)-biotin-AAA GGG CUG UUU UUC U[dT*]G CUG ACUUUC AGC CCC AAA-3)). This rC→dT* substitution was chosen to be the site of internal
fluorophore labeling for three reasons: (i) this nucleotide is one of the least conserved positionsin this region of the human TR pseudoknot motif (20), (ii) functional studies of the hTR PK haveshown that a complementary mutation of C106 did not affect in vitro activity of telomerase(13,49), and (iii) structural models of the minimal WT hTR PK depict this nucleotide as being
completely solvent exposed and devoid of hydrogen-bonding interactions with any of the!

184surrounding functional groups (20). The amino-modified nucleotide within strand 1 is reactedwith NHS-ester functionalized Cy5 dye (GE Healthcare) following the manufacturer’s suggested
protocol. Removal of unreacted Cy5 dyes is accomplished via micro centrifuge spin columns(Thermo Fisher Scientific, Inc). The second oligonucleotide, strand 2, contains a terminal 3) Cy3
attached via phosphoramidite chemistry: (5)-phosphate-CAA AAA AUG UCA GCA A-Cy3-3)).
Based on modifications to existing RNA ligation techniques (48,50), strands 1 and 2 areannealed together by heating to 85 °C at a 1:3 molar ratio in 50 mM hemisodium HEPES, 100mM NaCl, pH 7.5 and allowed to slowly cool to 37 °C. T4 RNA ligase I (New England BioLabs,
Inc.) is then added to the mixture following the manufacturer’s recommended procedures, withthe reaction held at 37 °C for 2 hours to allow for maximal ligation of the two strands (Figure7.1). The product is purified via HPLC using a reverse phase column (Agilent Technologies)
resulting in completely ligated RNA constructs containing Cy3, Cy5, and the biotin functionalgroup.
The mutant DKC hTR PK construct is created in a similar fashion, with the only
exception being that strand 1 for the DKC hTR PK contains the GC→AG mutation at positions
108-109 associated with the genetic disorder dyskeratosis congenita. A second wild-type
construct with alternative labeling locations (WTalt hTR PK) serves as a control to demonstratethat the observed conformational transitions are due to disruption and formation of the P3, rather
than P2, region of the pseudoknot (Supplemental Figure S7.1). This WTalt hTR PK construct issynthesized using the same procedures described above with two different oligonucleotides:(strand 1alt: 5)-Cy3-GGG CUG UUU UUC-3) and strand 2alt: 5)-phosphate-UCG CUG ACUUUC AGC CC[dT*] AAA CAA AAA AUG UCA GCA AAA-biotin-3)). These alternate
oligonucleotides shift the Cy 3 and Cy5 labeling sites from nucleotides 50 and 17 in the WT hTR!

185

PK to nucleotides 1 and 30 in the WTalt hTR PK (numbering from 5)-end of the smFRET
constructs used within this study).
7.5.2

Single-molecule Microscope
All single-molecule experiments are performed with an inverted confocal fluorescencemicroscope (Olympus Corporation) outfitted with a 1.2 NA water-immersion objective(Olympus Corporation)(Figure 7.9).

Figure 7.9 Single-molecule fluorescence microscope. Diagram of the experimental setup anddata collection (see Section 7.5.2 for details).
Direct excitation of the Cy3 (donor) fluorophore is accomplished via a pulsed (20 MHz) 532 nm
laser (Time Bandwidth Products Inc.). As needed, direct excitation of the Cy5 fluorophore(acceptor) can also be accomplished with a pulsed 635 nm laser diode (PicoQuant GmbH)
triggered with a fixed 25 ns delay with respect to the 532 nm donor excitation pulse. The lightfrom both excitation sources is coaxially aligned and directed into the back aperture of theobjective, resulting in spatially overlapped laser foci and the ability for Alternating Laser!

186

EXcitation methods (ALEX)(51-53) to verify presence of both donor and acceptor dyes on any
given single molecule construct.
Photons are collected by the same objective in epifluorescence and focused through aspatial pinhole (50 µm) aligned for optimal fluorescence collection from the laser excitation
volume. After passing through the pinhole, the light is spatially separated by polarization (i.e.,
perpendicular, parallel) and color (i.e., donor, acceptor) before being re-focused onto one of four
single-photon avalanche photo diodes (APD, PerkinElmer Inc.) coupled to a time-correlated
single-photon counting (TCSPC) module (Becker & Hickl GmbH) (Figure 7.9). The arrival ofeach fluorescence photon is recorded with respect the exciting laser pulse (microtime) and startof the experiment (macrotime). The stream of photon macrotimes is used to construct a donoracceptor fluorescence time trajectory, which is a visual representation of the raw data collectedin both the freely diffusing and surface immobilized smFRET experiments (see below).
Conversely, the photon microtimes are used to construct fluorescence lifetime profiles of the twodyes, which, among other things, can be used to ensure isotropic rotational diffusion of the donor
and acceptor fluorophores and thus the absence of any dye–RNA interactions.
7.5.3

Freely Diffusing smFRET Experiments
Freely diffusing experiments, similar to two-color coincidence detection one-color

excitation experiments (8,54), are performed at relatively high average laser powers (≈ 75 µW)
with low RNA concentrations (≈ 125 pM). At these concentrations there is, on average,
significantly fewer than one RNA molecule in the confocal excitation volume at any time (Figure
7.10a). As an individual RNA in solution stochastically passes through the excitation volume of
the focused laser beams, a short (< 1ms) burst of fluorescence is emitted that corresponds to theresonance time of the RNA in the confocal volume (Figure 7.10b). Sorting photon arrival times!

187

via ALEX methods provides the crucial ability to restrict analysis only to constructs containing
both non-photobleached donor and acceptor fluorophores. This is accomplished by assigning athreshold of 20 KHz on the minimum photon count rate for a given fluorescent burst to ensuresufficient determination of EFRET, with the reported values insensitive within quoted uncertainties
to changes in such a choice of threshold.

Figure 7.10 Freely diffusing experiments. a) Schematic representation of single-molecules freely
diffusing in and around the overlapping excitation volumes of the red and green lasers (seeMaterials and Methods for details). b) Freely diffusing data of the human telomerase RNA
pseudoknot obtained using ALEX methods, with the four possible outcomes for the fluorescentmolecules. ALEX filtering permits events from dually labeled donor-acceptor molecules to be
selectively considered for data analysis.

!

188

Data analysis is accomplished by calculating EFRET values for all ALEX sorted bursts
with both the donor and acceptor dyes present, which are then used to construct a histogram
representing equilibrium distribution of EFRET for the hTR PK molecules under a given set ofexperimental conditions(55). From the minimum (EFRET ≈ 0.55) in such a bimodal distribution(Figure 7.2), it is easy to determine which fluorescent bursts arise from molecules in the hairpin(unfolded, EFRET < 0.55) or pseudoknot (folded, EFRET > 0.55) conformation. Due to sample
purity and the use of ALEX methods, the number of high and low EFRET bursts is directly
proportional to the concentration of each of the conformers at equilibrium; thus the ratio of highand low EFRET burst frequencies provides direct experimental access to the equilibrium constant(Figure 7.1c).
7.5.4

Surface Immobilized smFRET Experiments
For surface immobilized experiments, molecules are non-covalently attached to thesurface via biotin-streptavidin chemistry, with a typical surface coverage of less than one
molecule per square micron. To generate an image of the surface immobilized molecules, apiezo-electric stage raster scans a 10 × 10 µm region of the sample with respect to themicroscope objective to generate a 256 × 256 pixel image. Each diffraction-limited feature in theimage corresponds to the fluorescence from a single surface-immobilized molecule. The stage isthen used to position the sample so that the laser light is focused at the center of an individual
fluorescence spot, from which the fluorescence can be continuously observed in real-time.
Again, the use of ALEX filtered surface images ensure that only dually labeled molecules are
considered for data analysis. As mentioned previously, the collected donor and acceptor photon
streams are used to construct both fluorescence and EFRET time trajectories, which for surfaceimmobilized experiments depict individual molecules actively switching between conformations!

189with distinct FRET efficiencies. Rate constants for these conformational transitions arecalculated by constructing cumulative distribution plots of the dwell times for the pseudoknot(EFRET > 0.55) and hairpin (EFRET < 0.55) conformations.
7.5.5

smFRET Temperature Control
For all of the temperature-dependent smFRET experiments, a heated objective collar(Bioptechs Inc.) and Peltier-base stage heater (Instec, Inc.) are used to provide thermal control ofthe sample with temperature gradients < 0.1 °C/cm. The absolute sample temperature isdetermined from measurements at the focus of the excitation source using a micro-thermocouple.
All other experiments are performed at room temperature (20-22 °C).
7.5.6

smFRET Buffer Conditions
Experiments are performed in a standard fluorescence-imaging buffer (50 mM HEPES,

25 mM KOH, 10 mM NaOH, 95 mM KCl, 2 mM TROLOX, pH 7.7), which consists of the wellcharacterized protocatechuic acid/protocatechuate-3,4-dioxygenase oxygen scavenging system(100 nM PCD and 5 mM PCA)(56). For all experiments, the molar concentrations of each of thebuffer components are held constant by accounting for changes in solution volume and density
due to the added solute.

!

190

References
1.

Wynford-Thomas, D.; D. Kipling. The End-Replication Problem, Nature, 1997, 389,
551-551.

2.

Blackburn, E. H. Switching and Signaling at the Telomere, Cell, 2001, 106, 661-673.

3.

Blackburn, E. H. The End of the (DNA) Line, Nat. Struct. Biol., 2000, 7, 847-850.

4.

Greider, C. W.; E. H. Blackburn. Tracking Telomerase, Cell, 2004, 116, S83-86.

5.

Cech, T. R. Beginning to Understand the End of the Chromosome, Cell, 2004, 116, 273279.

6.

Hengesbach, M.; B. M. Akiyama; M. D. Stone. Single-Molecule Analysis of Telomerase
Structure and Function, Curr. Opin. Chem. Biol., 2011, 15, 845-852.

7.

Mihalusova, M.; J. Y. Wu; X. Zhuang. Functional Importance of Telomerase Pseudoknot
Revealed by Single-Molecule Analysis, Proc. Natl. Acad. Sci. USA, 2011, 108, 2033920344.

8.

Yeoman, J. A.; A. Orte; B. Ashbridge; D. Klenerman; S. Balasubramanian. RNA
Conformation in Catalytically Active Human Telomerase, J. Am. Chem. Soc., 2010, 132,2852-2853.

9.

Wu, J. Y.; M. D. Stone; X. Zhuang. A Single-Molecule Assay for Telomerase StructureFunction Analysis, Nucleic Acids Res., 2010, 38, e16.

10.

Gavory, G.; M. F. Symmons; Y. Krishnan Ghosh; D. Klenerman; S. Balasubramanian.
Structural Analysis of the Catalytic Core of Human Telomerase RNA by FRET and
Molecular Modeling, Biochemistry, 2006, 45, 13304-13311.

11.

Collins, K. The Biogenesis and Regulation of Telomerase Holoenzymes, Nat. Rev. Mol.
Cell. Biol., 2006, 7, 484-494.

12.

Chen, J. L.; C. W. Greider. Telomerase RNA Structure and Function: Implications for
Dyskeratosis Congenita, Trends Biochem. Sci., 2004, 29, 183-192.

13.

Chen, J. L.; M. A. Blasco; C. W. Greider. Secondary Structure of Vertebrate Telomerase
RNA, Cell, 2000, 100, 503-514.

14.

Gilley, D.; E. H. Blackburn. The Telomerase RNA Pseudoknot is Critical for the Stable
Assembly of a Catalytically Active Ribonucleoprotein, Proc. Natl. Acad. Sci. USA, 1999,
96, 6621-6625.

15.

Tzfati, Y.; Z. Knight; J. Roy; E. H. Blackburn. A Novel Pseudoknot Element is Essential
for the Action of a Yeast Telomerase, Genes Dev., 2003, 17, 1779-1788.

!

191

16.

Reipa, V.; G. Niaura; D. H. Atha. Conformational Analysis of the Telomerase RNA
Pseudoknot Hairpin by Raman Spectroscopy, RNA, 2007, 13, 108-115.

17.

Moriarty, T. J.; D. T. Marie-Egyptienne; C. Autexier. Functional Organization of Repeat
Addition Processivity and DNA Synthesis Determinants in the Human Telomerase
Multimer, Mol. Cell. Biol., 2004, 24, 3720-3733.

18.

Autexier, C.; R. Pruzan; W. D. Funk; C. W. Greider. Reconstitution of Human
Telomerase Activity and Identification of a Minimal Functional Region of the Human
Telomerase RNA, Embo. J., 1996, 15, 5928-5935.

19.

Bachand, F.; I. Triki; C. Autexier. Human Telomerase RNA-Protein Interactions, Nucleic
Acids Res., 2001, 29, 3385-3393.

20.

Kim, N. K.; Q. Zhang; J. Zhou; C. A. Theimer; R. D. Peterson; J. Feigon. Solution
Structure and Dynamics of the Wild-Type Pseudoknot of Human Telomerase RNA, J.
Mol. Biol., 2008, 384, 1249-1261.

21.

Comolli, L. R.; I. Smirnov; L. Xu; E. H. Blackburn; T. L. James. A Molecular Switch
Underlies a Human Telomerase Disease, Proc. Natl. Acad. Sci. USA, 2002, 99, 1699817003.

22.

Theimer, C. A.; C. A. Blois; J. Feigon. Structure of the Human Telomerase RNA
Pseudoknot Reveals Conserved Tertiary Interactions Essential for Function, Mol. Cell.,
2005, 17, 671-682.

23.

Theimer, C. A.; L. D. Finger; L. Trantirek; J. Feigon. Mutations Linked to Dyskeratosis
Congenita Cause Changes in the Structural Equilibrium in Telomerase RNA, Proc. Natl.
Acad. Sci. USA, 2003, 100, 449-454.

24.

Theimer, C. A.; L. D. Finger; J. Feigon. YNMG Tetraloop Formation by a Dyskeratosis
Congenita Mutation in Human Telomerase RNA, RNA, 2003, 9, 1446-1455.

25.

Chen, G.; J. D. Wen; I. Tinoco, Jr. Single-Molecule Mechanical Unfolding and Foldingof a Pseudoknot in Human Telomerase RNA, RNA, 2007, 13, 2175-2188.

26.

Hengesbach, M.; N. K. Kim; J. Feigon; M. D. Stone. Single-Molecule FRET Reveals the
Folding Dynamics of the Human Telomerase RNA Pseudoknot Domain, Angew. Chem.
Int. Ed. Engl., 2012, 51, 5876-5879.

27.

Cao, S.; S. J. Chen. Biphasic Folding Kinetics of RNA Pseudoknots and Telomerase
RNA Activity, J. Mol. Biol., 2007, 367, 909-924.

28.

Cao, S.; S. J. Chen. Predicting RNA Pseudoknot Folding Thermodynamics, Nucleic
Acids Res., 2006, 34, 2634-2652.

29.

Li, P. T.; J. Vieregg; I. Tinoco, Jr. How RNA Unfolds and Refolds, Annu. Rev. Biochem.,
2008, 77, 77-100.

!

192

30.

Bokinsky, G.; X. Zhuang. Single-Molecule RNA Folding, Acc. Chem. Res., 2005, 38,
566-573.

31.

Roy, R.; S. Hohng; T. Ha. A Practical Guide to Single-Molecule FRET, Nat. Methods,
2008, 5, 507-516.

32.

Zhao, R.; D. Rueda. RNA Folding Dynamics by Single-Molecule Fluorescence
Resonance Energy Transfer, Methods, 2009, 49, 112-117.

33.

Aleman, E. A.; R. Lamichhane; D. Rueda. Exploring RNA Folding One Molecule at aTime, Curr. Opin. Chem. Biol., 2008, 12, 647-654.

34.

Holmstrom, E. D.; J. L. Fiore; D. J. Nesbitt. Thermodynamic Origins of Monovalent
Facilitated RNA Folding, Biochemistry, 2012, 51, 3732-3743.

35.

Fiore, J. L.; E. D. Holmstrom; D. J. Nesbitt. Entropic Origin of Mg2+-Facilitated RNA
Folding, Proc. Natl. Acad. Sci. USA, 2012, 109, 2902-2907.

36.

Bartley, L. E.; X. W. Zhuang; R. Das; S. Chu; D. Herschlag. Exploration of theTransition State for Tertiary Structure Formation Between an RNA Helix and a Large
Structured RNA, J. Mol. Biol., 2003, 328, 1011–1026.

37.

Fiore, J. L.; B. Kraemer; F. Koberling; R. Erdmann; D. J. Nesbitt. Enthalpy-Driven RNA
Folding: Single-Molecule Thermodynamics of Tetraloop–Receptor Tertiary Interaction,
Biochemistry, 2009, 48, 2550–2558.

38.

Shelton, V. M.; T. R. Sosnick; T. Pan. Applicability of Urea in the Thermodynamic
Analysis of Secondary and Tertiary RNA Folding, Biochemistry, 1999, 38, 16831-16839.

39.

Lambert, D.; D. E. Draper. Effects of Osmolytes on RNA Secondary and Tertiary
Structure Stabilities and RNA-Mg2+ Interactions, J. Mol. Biol., 2007, 370, 993-1005.

40.

Fersht, A. R.; A. Matouschek; L. Serrano. The Folding of an Enzyme .1. Theory ofProtein Engineering Analysis of Stability and Pathway of Protein Folding, J. Mol. Biol.,
1992, 224, 771-782.

41.

Priyakumar, U. D.; C. Hyeon; D. Thirumalai; A. D. MacKerell. Urea Destabilizes RNA
by Forming Stacking Interactions and Multiple Hydrogen Bonds with Nucleic Acid
Bases, J. Am. Chem. Soc., 2009, 131, 17759-17761.

42.

Lambert, D.; D. E. Draper. Denaturation of RNA secondary and tertiary structure byurea: simple unfolded state models and free energy parameters account for measured mvalues, Biochemistry, 2012, 51, 9014-9026.

43.

Williams, S.; T. P. Causgrove; R. Gilmanshin; K. S. Fang; R. H. Callender; W. H.
Woodruff; R. B. Dyer. Fast Events in Protein Folding: Helix Melting and Formation in a
Small Peptide, Biochemistry, 1996, 35, 691-697.

!

193

44.

Yingling, Y. G.; B. A. Shapiro. The Impact of Dyskeratosis Congenita Mutations on the
Structure and Dynamics of the Human Telomerase RNA Pseudoknot Domain, J. Biomol.
Struct. Dyn., 2007, 24, 303-319.

45.

Akiyama, B. M.; M. D. Stone. Assembly of Complex RNAs by Splinted Ligation,
Methods Enzymol., 2009, 469, 27-46.

46.

Moore, M. J.; C. C. Query. Joining of RNAs by Splinted Ligation, Method Enzymol.,
2000, 317, 109-123.

47.

Stark, M. R.; J. A. Pleiss; M. Deras; S. A. Scaringe; S. D. Rader. An RNA LigaseMediated Method for the Efficient Creation of Large, Synthetic RNAs, RNA, 2006, 12,2014-2019.

48.

Moore, M. J., Query, C.C. Uses of Site-Specifically Modified RNAs Constructed by
RNA Ligation, In RNA:Protein Interactions: A Practical Approach (Smith, C. W. J.,
Ed.), pp 75-140, Oxford University Press Inc., New York.1998

49.

Chen, J. L.; C. W. Greider. Functional Analysis of the Pseudoknot Structure in Human
Telomerase RNA, Proc. Natl. Acad. Sci. USA, 2005, 102, 8080-8085; discussion 80778089.

50.

Lemay, J. F.; J. C. Penedo; R. Tremblay; D. M. Lilley; D. A. Lafontaine. Folding of the
Adenine Riboswitch, Chem. Biol., 2006, 13, 857-868.

51.

Lee, N. K.; A. N. Kapanidis; Y. Wang; X. Michalet; J. Mukhopadhyay; R. H. Ebright; S.
Weiss. Accurate FRET Measurements within Single Diffusing Biomolecules using
Alternating-Laser Excitation, Biophys. J., 2005, 88, 2939-2953.

52.

Kapanidis, A. N.; N. K. Lee; T. A. Laurence; S. Doose; E. Margeat; S. Weiss.
Fluorescence-Aided Molecule Sorting: Analysis of Structure and Interactions by
Alternating-Laser Excitation of Single Molecules, Proc. Natl. Acad. Sci. USA, 2004, 101,
8936–8941.

53.

Kapanidis, A. N.; T. A. Laurence; N. K. Lee; E. Margeat; X. X. Kong; S. Weiss.
Alternating-Laser Excitation of Single Molecules, Acc. Chem. Res., 2005, 38, 523-533.

54.

Orte, A.; R. Clarke; S. Balasubramanian; D. Klenerman. Determination of the Fractionand Stoichiometry of Femtomolar Levels of Biomolecular Complexes in an Excess of
Monomer using Single-M olecule, Two-Color Coincidence Detection, Anal. Chem.,
2006, 78, 7707-7715.

55.

Fiore, J. L.; J. H. Hodak; O. Piestert; C. D. Downey; D. J. Nesbitt. Monovalent andDivalent Promoted GAAA-Tetraloop–Receptor Tertiary Interactions from Freely
Diffusing Single-Molecule Studies, Biophys. J., 2008, 95, 3892-3905.

!

194

56.

!

Aitken, C. E.; R. A. Marshall; J. D. Puglisi. An Oxygen Scavenging System for
Improvement of Dye Stability in Single-Molecule Fluorescence Experiments, Biophys. J.,
2008, 94, 1826-1835.

195

Chapter 8

Vitamin B12-facilitates formation of a regulatory RNA kissing loop*

8.1

Introduction
Riboswitches are noncoding structural elements in the leader sequence of certain

bacterial messenger RNAs that regulate downstream gene expression in a ligand-dependent
fashion(1). The first experimentally verified riboswitch was found in mRNAs that code for the
expression of a cobalamin (vitamin B12) transport protein (i.e., btuB in E. coli)(2,3). These
structural elements function to maintain intracellular concentrations of their cognate ligand (i.e.,cobalamin), which is a critical cofactor for methyltransferase and isomerase reactions involved in
S-adenosylmethionine and succinyl-CoA biosynthesis, respectively(4).The cobalamin clan ofriboswitches (Rfam: CL00101) is one of the most abundant and ubiquitous cis-acting mRNA
regulatory elements throughout the bacterial domain of life(5). The importance of these RNAs
and their associated effector molecule is further highlighted by the fact that only certain speciesof Archaea and Bacteria can biosynthesize cobalamin(6,7), and yet it is essential to manyorganisms, most notably humans(8).
Generally, all riboswitches consist of two interacting domains that, together, result inbiological functionality. Specifically, ligand binding in the receptor (or aptamer) domain
promotes a conformational change in the regulatory domain (or expression platform), which
modulates the production of a downstream gene(9,10). Single-molecule fluorescence resonanceenergy transfer (smFRET) is ideally suited(11) and frequently used to observe these ligand-*

To be submitted196

induced conformational transitions(12-17). However, many of these investigations are unable tomonitor conformational transitions associated with the regulatory domain, and, until now, noneof them have been able to unambiguously link: (i) ligand binding to (ii) conformationaltransitions within a structural switch. Here we use these smFRET techniques to independently
and simultaneously observe these two processes.
A recent study reported the first x-ray crystal structure of an entirely biologicallyfunctional cobalamin riboswitch containing both the receptor and regulatory domains(18). This
crystallographic model revealed that the recognition and regulatory domains communicate via atertiary interaction in order to confer biological functionality. Additionally, cell-based
experiments conducted on these RNAs demonstrate that this tertiary interaction modulates gene
expression in a vitamin B12-dependent fashion(18). Specifically, hydroxocobalamin, a variant ofvitamin B12, was shown to facilitate formation of a kissing loop (KL) interaction, resulting insequestration of the ribosome binding site (RBS). Although these structural and cell-based
experiments have unveiled information about the mechanism of action associated with thisriboswitch, many detailed questions regarding both ligand binding kinetics and conformational
dynamics remain unanswered.
To this end, a combination of fluorescence techniques—smFRET and cell-based
fluorescent-reporter assays—has been used to offer the first, to our knowledge, detailed
mechanistic account of a biologically functional riboswitch based on a collection of both cellularand smFRET data. The primary sequence (GenBank accession: AACY021350931.1/557-456) of
the env8 hydroxocobalamin (env8HyCbl, Figure 8.1a) riboswitch used throughout theseexperiments comes from a comparative genomics analysis of structured noncoding RNAs(19). It
is member of the AdoCbl-variant family (Rfam: RF01689), which is one of three members

197within the larger cobalamin riboswitch clan (Rfam: CL00101)(20). Previous ligand binding
experiments have revealed that env8HyCbl selectively binds vitamin B12 cofactors with smaller
β-axial ligands (e.g., hydroxo- or methylcobalamin) as opposed to the bulkier adenosylcobalamincofactor(18). Synthetic modifications to this RNA (Figure 8.1a) allow smFRET-based
experiments to monitor, in real-time and under equilibrium, the docking/undocking of the L5L13 regulatory KL in the presence or absence of ligand (Figure 8.1b).

Figure 8.1. Schematic representation of the a) secondary and b) tertiary structure of theenv8HyCbl riboswitch, complete with the Cy3 (green star), Cy5 (red star), and biotin (bluerectangle) synthetic moieties. The loops, L5 (red) and L13 (green) form a regulatory RNA
kissing-loop (KL), which contains the purine-rich ribosome binding site (shadowed green
nucleotides). Binding of hydroxocobalamin (bronze spheres, HyCbl) facilitates formation of theKL, which decreases the distance between the Cy3 (donor) and Cy5 (acceptor) fluorophores
resulting in more efficient energy transfer (EFRET).

198

Simultaneously, hydroxocobalamin-dependent quenching(21) of Cy3 is used as an orthogonal
experimental observable to monitor the kinetics of ligand binding independently of
conformation. The results of these smFRET experiments provide substantial support for a fourstate kinetic model that highlights several key biological features associated with this regulatory
switch (Figure 8.2): (i) KL docking/undocking occurs in the absence of ligand and is dependent
on Mg2+, (ii) ligand binding only occurs when the KL is undocked and (iii) KL
docking/undocking also occurs in the presence of ligand and is dependent on Mg2+.

Figure 8.2 Four-state kinetic model for the env8HyCbl riboswitch represented a) graphically andb) symbolically. The four macroscopic conformations are linked by three coupled equilibria: (i)KL docking in the absence of ligand, (ii) ligand binding in the undocked conformation, and (iii)KL docking in the presence of ligand. Formation of the KL decreases the inter-dye distance,
resulting in more efficient fluorescence resonance energy transfer (EFRET). Ligand binding
quenches Cy3, which decreases the total fluorescence of Cy3 and Cy5 and can be monitored
independent of KL docking/undocking.
Additionally, the single-molecule results reveal that, in the presence of Mg2+, ligand binding
preferentially stabilizes the docked conformation. Finally, results from cell-based reporter assays
are used to relate these biophysical findings with the cellular function of this RNA. The highly199correlated experimental observations reveals that formation of the regulatory switch (i.e., L5-L13
KL) is responsible for repression of gene expression, regardless of whether or not the ligand isbound to the RNA. As a whole, these findings expose a great deal of information about the
dynamical relationship between structure and function in cobalamin riboswitches, which can beapplied to various other functional RNAs.
8.2

Results
Both freely diffusing and surface-immobilized experiments (See Experimental Methods)

have been used to characterize the conformational transitions associated with the regulatory KLof the env8HyCbl riboswitch. Briefly, freely diffusing smFRET experiments make use of theextremely small volume associated with the diffraction-limited focus of a high numericalaperture objective. At low concentrations of Cy3-Cy5 labeled RNA (≈ 100 pM), this volume isvoid of fluorophores ≈98% of the time. Occasionally, an individual molecule will stochasticallydiffuse through this volume, resulting in a burst of fluorescence limited in duration by the
diffusion constant of the RNA (≈ 1 ms). The efficiency of fluorescence resonance energy transfer(EFRET) can be calculated for thousands of fluorescent bursts and compiled into a histogram todescribe the probability of observing any given EFRET value, which is related to the conformationof the RNA.
Alternatively, individual Cy3-Cy5 labeled RNAs can be immobilized to a microscope
coverslip using biotin-streptavidin chemistry, which prevents the molecule from freely diffusing
in solution. The diffraction-limited focus of the confocal fluorescence microscope can be used tocontinuously excite the fluorophores and subsequently detect fluorescence. The collected signal
is used to determine the EFRET of a single RNA as a function of time, where changes in EFRET
result from discrete conformations transitions.

200

The two single-molecule fluorescence experiments described above are used to rigorously
characterize the in vitro behavior of the env8HyCbl riboswitch. Complementary cell-based
fluorescence reporter assays are used to compare and contrast the in vitro and cellular behaviorof the RNA. Specifically, these reporter assays are designed to monitor the regulation of green
fluorescent protein (GFP) expression provided by the env8HyCbl riboswitch.
8.2.1

KL docking equilibrium in the absence of ligand
As a first step, freely diffusing smFRET experiments have been used to explore theequilibrium behavior of the L5-L13 kissing loop (KL) interaction from the env8HyCbl
riboswitch in solutions free of ligand. Under standard buffer conditions (see Experimental
Methods), the distribution of EFRET values associated with thousands of individual fluorescently
labeled RNA constructs reveals (Figure 8.3a) that the riboswitch resides entirely in a single low
EFRET = 0.15(3) population. However, addition of MgCl2 is sufficient to promote KL docking(Figure 8.3a), as indicated by the Mg2+-dependent increase in the relative probability ofobserving molecules in a second, higher EFRET = 0.58(3) population. This high EFRET value
corresponds to an inter-dye distance of ≈ 50(3) Å and, given the experimental uncertainty in themeasurement, is consistent with the ≈ 47 Å prediction based on the recent crystal structure(18).
Accordingly, we attribute this high EFRET value to molecules residing in the docked KL
conformation (Dfree), with the low EFRET value corresponding to the undocked species (Ufree),where the subscript “free” denotes that the ligand is not bound the riboswitch (Figure 8.2b). The
necessity of Mg2+ for the formation of this interaction is supported by the x-ray crystal structure(PDB: 4FRN), which depicts divalent cations near the L5 and L13 loops(18). This is a notablefeature among other KL x-ray crystal structures(22,23) and likely reflects the importance of
Mg2+ in the formation of these types of tertiary interactions.

201

Figure 8.3 Freely diffusing smFRET experiments depict the Mg2+-dependence of the KL
docking equilibrium in the a) absence and b) presence of 1000 nM HyCbl. c) The presence ofHyCbl (bronze circles) decreases the concentration of divalent cations (i.e., [Mg2+]½) required
for FD = 0.5.
The residual probability associated with observing the low EFRET population at high[Mg2+] results from molecules that are unable to form the L5-L13 KL interaction (≈ 15 %), as
determined in the surface immobilized experiments described later (Section 8.2.5). The
normalized probability of observing the two EFRET distributions, when corrected for a 15 %nonresponsive population, can be used to characterize the equilibrium behavior of the riboswitch.
Specifically, the fractional occupancy of molecules in the docked or undocked conformation canbe calculated using Eq. 8.1a and 1b, respectively.

202

!"!"## ! = ! ! !
!"!"## ! = ! ! !

! !!"##!"## !!!! !!"##! !!"##!"## !!!! !!"##(Eq. 8.1a)(Eq. 8.1b)

The parameters P(Dfree) and P(Ufree) describe the probability of observing the docked andundocked conformations, respectively. Again, the subscript “free” is used to indicate that theriboswitches are not bound to ligands. The Mg2+ concentration required to achieve FD = 0.5 (i.e.,[Mg2+]½) is obtained by fitting the data to a standard Hill-type binding model. For data collectedin the absence of HyCbl, such an analysis results in a [Mg2+]½ = 2.2(2) mM (Figure 8.3c), which
is nearly twice normal physiological concentrations of free Mg2+(24). Although the regulatory
KL can form in the absence of HyCbl, the FDfree is only ≈ 0.22 in solutions that appropriately
mimic the ionic environment associated with the inside of a cell (e.g., 1 mM Mg2+, 125 mM K+,
10 mM Na+), suggesting that undocked conformation is more stable under these conditions.
To confirm that the high EFRET population results from formation of the L5-L13
regulatory switch, a second, doubly labeled RNA construct has also been studied. This construct,referred to as the L5-mutant env8HyCbl riboswitch (Supplemental Figure S8.1), contains adinucleotide substitution at positions 48 and 49 in L5 (underlined nucleotides, 5"-UACUUG-"3→ 5"-UACAAG-"3). The UU to AA mutation prevents formation of two Watson-Crick basepairs that are otherwise present in the wild-type construct and preferentially destabilizes thedocked conformation of the KL. As expected, this smFRET construct remains in the undocked(low EFRET) conformation even in solutions containing 30 mM MgCl2 (Supplemental FigureS8.1), thus confirming that the donor and acceptor fluorophores accurately report on theformation of this regulatory KL interaction.

203

Supplemental Figure S8.1. Results of freely diffusing experiments conducted on the L5-mutant
env8HyCbl construct. The L5-mutation (inset, underlined nucleotides) inhibits formation of the
KL interaction, which prevents occlusion of the ribosome binding site (inset, shadowed
nucleotides).
8.2.2

KL docking equilibrium in the presence of ligand
Analogous freely diffusing smFRET experiments have been conducted in the presence ofligand in order to interrogate the equilibrium behavior of this interaction in solutions where the
HyCbl concentration is ≈ 50 times larger than a previously reported(18) binding affinity of 19(4)nM. Without any Mg2+ in solution, fluorescent molecules reside entirely in a single low EFRET =0.12(3) population (Figure 8.3b) that is experimentally indistinguishable from the low EFRET =0.15(3) population observed in the absence of ligand (Figure 8.3a). As was the case before, Mg2+
is still required for the formation of the KL interactions, even under saturating concentrations of
HyCbl. Specifically, addition of MgCl2 into solutions containing 1000 nM HyCbl increases theprobability of observing a second, higher EFRET = 0.54(4) population (Figure 8.3b) that is also
experimentally indistinguishable from the high EFRET = 0.58(3) population noted previously inthe absence of ligand (Figure 8.3a).

204

Under these saturating HyCbl conditions, presumably every functional riboswitch isbound to a ligand. Therefore, the above observations support the existence of L5-L13 KL
equilibrium associated with the docked (DHyCbl) and undocked (UHyCbl) conformations of theRNA, where the “HyCbl” subscript denotes the presence of a bound ligand (Figure 8.2b).
Notably, the presence of a bound ligand reduces [Mg2+]½ by nearly an order of magnitude(Figure 8.3c), which indicates that under physiological free salt conditions (i.e., 1 mM Mg2+,
125 mM K+, 10 mM Na+) these ligand bound riboswitches reside predominantly in the docked
conformation (FDHyCbl ≈ 0.75). In conjunction with the results of previous experiments
performed in the absence of ligand, it is immediately obvious that HyCbl binding increases thestability of the docked conformation in solutions containing Mg2+(e.g., at 1 mM Mg2+ FDfree ≈0.22 < FDHyCbl ≈ 0.75). Moreover, the increased occupancy of the docked conformation, where
the L5-L13 KL is form, clearly demonstrates that ligand binding directly modulates theavailability of the ribosome binding site nucleotides in L13 of this regulatory switch.
8.2.3

Ligand binding properties
To directly explore the ligand-binding process, we exploit a unique photophysicalproperty of hydroxocobalamin (HyCbl). Specifically, the absorbance spectra of vitamin B12
cofactors, including HyCbl, overlap with the fluorescence spectra of yellow-green fluorescentdyes like Cy3, which allows this cofactor to effectively compete with the Cy5 for energy
transfer. Because HyCbl is a non-fluorescent acceptor, energy transfer to it results in quenched
fluorescence(21). We can utilize this quenching phenomenon to monitor the ligand binding
kinetics of the env8HyCbl riboswitch via time-dependent ensemble fluorometry. When HyCbl is
dissociated from the RNA, the average intermolecular distance between it (acceptor) and Cy3(donor) is large enough to neglect the effects of this quenching energy transfer process (Figure205

8.4a). However, upon ligand binding, the intermolecular distance between the two is reduced to≈ 23 Å, allowing for predominant energy transfer from the excited donor molecule (Cy3) to thenon-fluorescent acceptor (HyCbl).

Figure 8.4 Time-dependent ensemble fluorometry. a) HyCbl binding quenches the fluorescenceof Cy3 (see also Supplementary Figure S8.2). b) Elevated concentrations of ligand increases therate constant for establishing equilibrium. c) A linear-fit to the [HyCbl]-dependence of this rate
constant is used to measure the apparent ligand binding (appkbind) and dissociation (appkdiss) rate
constants.
Thus, energy transfer to HyCbl results in an additional non-radiative (knrad) component of the
Cy3 fluorescence lifetime (τfluor = 1/ (krad + knrad)), which leads to a decreased quantum yield(Q.Y. = krad / (krad + knrad)). As an attempt to quantify this ligand-dependent quenching, we havemeasured the fluorescence decay profiles of individual Cy3-only env8HyCbl RNAs in the206

presence and absence of 1000 nM HyCbl (Supplemental Figure S8.2) using the time-correlated
single-photon counting capabilities associated with the experimental apparatus (see Experimental
Methods).

Supplemental Figure S8.2 The fluorescence lifetime (τfluor) of Cy 3 as determined fromsurface-immobilized riboswitches bound to (light green triangles) or free of (dark green circles)
hydroxocobalamin (HyCbl).
These lifetime measurements (i.e., Cy3-τfluor(free) = 1.40(2) ns and Cy3-τfluor(HyCbl) =0.51(6) ns) effectively demonstrate the non-radiative quenching phenomenon associated withenergy transfer from Cy3 to HyCbl. It is important to stress that the presence of bound ligand
introduced a non-radiative decay process from the excited state that competes equally with bothradiative emission from the excited donor (i.e., Cy3 fluorescence) and energy transfer from theexcited donor to the acceptor (Cy5 fluorescence). Provided that there is sufficient signal-to-noise,
this will not alter the observed EFRET values associated with the docked and undocked
populations, as is demonstrated above (Section 8.2.2).
Time-dependent ensemble fluorescence experiments are used to monitor the HyCblinduced reduction in fluorescence intensity of RNA constructs labeled with Cy3 after addition ofexcess ligand ([HyCbl]/[RNA] > 10). The temporal decay of fluorescence intensity (Figure 8.4b)
maps out the rate constant responsible for establishing a binding equilibrium (keq). For a standard
207

bimolecular process, this rate constant (keq) can simply be described as a function of ligand
concentration using the ligand-binding (kbind) and dissociation (kdiss) rate constants (Eq. 8.2).
!!" = !!"## + !"#$%& !!"#$(Eq. 8.2)

However, in the present system, the rate constants only represent apparent parameters because
the ligand binding equilibrium is directly coupled to the two KL docking equilibria (Figure 8.2).
These two docking equilibria determine the fraction of undocked molecules that can either bindto (FUfree) or dissociate from HyCbl (FUHyCbl). The ligand dependence of appkeq is evident in
Figure 8.4c and well described by a straight line. The slope of such a linear fit reveals that appkbind= 1.0(1)×106 M-1 s-1, where as the intercept shows that appkdiss = 0.005(3) s-1. The ratio of thesetwo values describes the apparent HyCbl dissociation equilibrium constant (appKd = 5(3) nM).
These apparent kinetic parameters can be decoupled from the two KL docking equilibria bymaking a well-justified assumption, which is experimentally validated later in surfaceimmobilized kinetic studies of the KL docking equilibria (Section 8.2.5 and 8.2.6). Provided thedocking kinetics associated with the KL equilibria are much faster than the kinetics of the ligand
binding process, the apparent rate constants equal the true kbind and kdiss values scaled by thefractional populations of the binding competent (FUfree) and dissociation competent (FUHyCbl)
species, respectively (see Eq. 8.3a,b).
!!"#$ !"!"## = ! !""!!"#$(Eq. 8.3a)!!"## !"!"#$% = ! !""!!"##(Eq. 8.3b)

The values of FUfree ≈ 0.78 and FUHyCbl ≈ 0.25 are obtained from the freely diffusing data
depicted in Figure 8.3c at1 mM MgCl2. These values are used to calculate the true rate andequilibrium constants—denoted by a lack of the superscript “app”—that describe ligand binding208

by the env8HyCbl riboswitch at 1 mM MgCl2 (kbind = 1.3(2)×106 M-1 s-1, kdiss = 0.02(1) s-1, andKd, = 15(8) nM).
8.2.4

CBL-dependence of the KL docking equilibrium
The data sets described in the three previous sections (8.2.1-8.2.3) quantitatively

characterize all of the equilibrium constants required to predict the steady-state fractional
occupancy of the docked conformation as a function of [HyCbl] at any given [MgCl2]. These
predictions are valid only if the true HyCbl dissociation constant (Kd) is not directly influenced
by MgCl2. As an aside, many nucleic acid-ligand interactions are known to be highly sensitive to
Mg2+; however, the ligand binding process for the env8HyCbl riboswitch only seems to beindirectly influenced by Mg2+ via the two KL docking equilibria, as described in the following
section.
To experimentally test the steady-state model predictions, single-molecule freely
diffusing FRET methods are used to determine the fractional occupancy of the docked
conformation (FD) as HyCbl is titrated into solutions at five different fixed MgCl2 concentrations(0.0, 0.5, 1.0, 2.5, 20. mM). In solutions lacking Mg2+, the addition of ligand is unable topromote formation of the docked conformation (Figure 8.5a), as Mg2+ is required for theformation of the regulatory KL interaction between L5 and L13. At intermediate concentrationsof MgCl2 (e.g., 0.5, 1.0, and 2.5 mM), titration of HyCbl into solution does facilitate formationof the docked conformation (Figure 8.5b). This observation is certainly consistent with the fourstate model (Figure 8.2), wherein bound HyCbl, increases the stability of the regulatory KLinteraction (i.e., FDfree < FDHyCbl). Titration of HyCbl into solutions with saturating
concentrations of Mg2+ (e.g., 20 mM >> [Mg2+]½ ) reveals that ligand is unable to significantly
increase the FD (Figure 8.5c), which nicely demonstrates that Mg2+ alone is sufficient to209promote complete formation of the regulatory interaction responsible for sequestering theribosome binding site.

Figure 8.5 Freely diffusing single-molecule burst titrations of hydroxocobalamin (HyCbl) at
various concentrations of MgCl2. a) At 0 mM Mg2+, addition of ligand does not significantly
influence distribution of FRET values associated with the L5-L13 regulatory switch. b) Atintermediate concentrations of Mg2+(e.g., 0.5, 1.0, and 2.5 mM), addition of HyCbl doessignificantly alter the FRET distributions. c) 20 mM Mg2+ is sufficient to completely form theL5-L13 KL interaction, irrespective of [HyCbl].
Figure 8.6 compares the results of the single-molecule experiments with the model predictions(colored circles and dashed lines, respectively). This comparison demonstrates the remarkable
quantitative accuracy of the four state kinetic model (Figure 8.2) used to describe theconformational transitions associated with this biologically functional riboswitch. Most
importantly, this astonishing predictive power is a testament to the validity of the kinetic model,
including the Mg2+ independence of true ligand binding process (Kd).

210

Figure 8.6. Experimental validation of the four-state kinetic model for the env8HyCbl
riboswitch. The experimentally determined fractional occupancy of the docked conformation(FD, colored circles) is well described by the steady-state solution (dotted line) to the four-statemodel (Figure 8.2) over a wide range of Mg2+ and HyCbl concentrations.
8.2.5

KL docking kinetics in the absence of ligand
To exploit the capabilities of our single-molecule microscopy apparatus for probing

kinetics, fluorescence trajectories of individual surface-immobilized molecules have beenrecorded to observe the KL docking dynamics in real time (Figure 8.7a-c). In the absence ofHyCbl (Figure 8.7a), single RNAs display strongly anti-correlated fluctuations in donor andacceptor fluorescence, resulting from rapid transitions between the two well-separated FRET
states. These FRET values are in excellent agreement with those measured in the freely diffusing
experiments, which supports the conclusion that these time trajectories depict single-molecules
transitioning between the docked (Dfree) and undocked (Ufree) conformations associated with theL5-L13 KL interaction. Kinetic analysis of the surface-immobilized data yields hundreds ofindividual dwell times that are compiled together and fit to single-exponential decays in order toyield the rate constants associated with docking and undocking. These rate constants reveal that,in ligand free solutions at 1 mM Mg2+, kdock,free = 1.2(3) s-1 and is slower than kundock,free = 6(2)s-1,
resulting in FDfree = 0.17(7). A titration with respect to Mg2+, analogous to the ones from the211freely diffusing smFRET experiments, reveals that increasing divalent cation concentration
accelerates the docking and slows the undocking rate constants, respectively.

Figure 8.7. Kissing-loop docking kinetics in the absence of bound ligand. Representative a)
fluorescence and b) FRET time-trace from a surface immobilized env8HyCbl riboswitch. c) Rate
constants are determined using both a dwell time analysis and a 2-state Bayesian maximumlikelihood model (orange line). d) Kinetic parameters are used to calculate the fractional
occupancy of the docked conformation in the absence of HyCbl (FDfree), which reveals that thesurface immobilized results are in accordance with the freely diffusing experiments.
Both of which contribute nearly equally to Mg2+-dependent formation of the regulatory KL(Figure 8.7c). As alluded to above, the fact that individual surface-immobilized molecules dockcompletely (FDfree ≈ 1.0) at high [MgCl2] confirms that the residual probability (≈ 15%) for
observing the low EFRET population results from inactive molecules that cannot form the L5-L13
interaction. It is important to emphasize that the results for both freely diffusing and surface
immobilized studies are in complete agreement as indicated by the circles and stars in Figure212

8.7d, respectively. Such a plot convincingly demonstrates that surface immobilization does not
influence the biophysical behavior of this RNA.
8.2.6

KL docking kinetics in the presence of ligand
To complete the single-molecule kinetic characterization of this riboswitch, fluorescencetime-trajectories of individual surface-immobilized molecules are recorded in the solutions
containing 2.5 nM HyCbl and 1 mM Mg2+ (Figure 8.8). These results further support the
proposed kinetic model (Figure 8.2) for ligand-facilitated formation of the regulatory switch.
Specifically, the time-trajectories in the presence of HyCbl display distinct regions with vastlydifferent fluorescence intensities. Some regions exhibit normal fluorescence signals for both thedonor and the acceptor fluorophores, which are well resolved and anti-correlated (Figure 8.8b).
In the other, intervening regions, the total fluorescence signal is significantly quenched (Figure8.8c). The rate constants associated with the anti-correlated fluorescence fluctuations in thenormal regions mimic those from molecules studied in the absence of hydroxocobalamin (Figure
8.8b vs. 8.7b), as would be expected when the ligand is not bound to the RNA. Conversely, thequenched regions result from ligand binding events that localize HyCbl near Cy3, which
introduces additional nonradiative relaxation pathways. Despite the substantially reduced signal
in these regions, subtle fluctuations in acceptor intensity that exceed background noise areclearly observable and can be attributed to docking and undocking of the KL interaction during
the long ligand-bound episodes. As an attempt to quantitatively measure the docked andundocked dwell times in the ligand-bound state, a Bayesian maximum-likelihood model(25) isused to analyze the two-state fluctuations evident in the quenched regions of the fluorescence
trajectories (Figure 8.8c). The results of such an analysis reveal that, when the ligand is bound to
the env8HyCbl riboswitch in solution containing 1 mM MgCl2, the docking rate only decreases

213

two fold (kdock,HyCbl = 0.64(9) vs. kdock,free = 1.2(3) s-1), but that the undocking rate is much moresensitive and decreases significantly by a ≈ 23-fold (kundock,HyCbl = 0.26(8) vs. kundock,free = 6(2)s-1).

Figure 8.8 Surface immobilized KL docking kinetics in the presence of hydroxocobalamin(HyCbl). a) Representative fluorescence trajectory from a surface immobilized molecule in thepresence of 2.5 nM HyCbl. While in the undocked conformation, ligand binding (arrow)significantly diminishes the fluorescence intensity, which returns to normal after dissociation. b)
When the ligand is not bound, the docking kinetics of the regulatory KL are experimentallyindistinguishable from those in the absence of any HyCbl (Figure 8.7b). c) When the ligand isbound, a maximum-likelihood 2-state model (black line) is used to quantify the dwell times
associated with the subtle fluctuations in acceptor fluorescence resulting from KL docking andundocking. A comparison between the two KL docking equilibria reveals that HyCbl binding
significantly decreases the undocking rate constant (kundock).
The HyCbl-induced change to the KL undocking kinetics give rise to substantially morefavorable formation of the L5-L13 regulatory switch (FDfree = 0.17(7) vs. FDHyCbl = 0.7(1)), as
also observed in the original freely diffusing experiments (FDfree ≈ 0.22 vs. FDHyCbl ≈ 0.75).

214

Supplemental Figure S8.3. Representative surface immobilized trajectories revealing the
HyCbl-induced decrease in fluorescence intensity of both Cy3 (green squares) and Cy5 (pinkcircles). Inspection of trajectories reveals that HyCbl binding events (black arrows) can only
occur when the RNA is in the undocked (low EFRET) conformation. This is consistent with theassumptions of our 4-state kinetic model, whereby formation of the L5-L13 KL interactions
prevents HyCbl binding/dissociation events. Only trajectories that exhibit clean fluctuations in
acceptor fluorescence after a decrease in total fluorescence are considered for analysis. This
constraint excludes molecules that have undergone other photophysical processes resulting in adecrease in fluorescence (e.g., blinking or bleaching).

215

Further inspection of the fluorescence trajectories reveals that: (i) HyCbl can only bind andquench fluorescence when the RNA is in the undocked (low EFRET) conformation (Supplemental
Figure S8.3) and (ii) ligand-bound dwell times can be longer than a hundred seconds (Figure8.8a). These bound-state lifetimes are consistent with those predicted from the ensemble timedependent fluorescence experiments (Figure 8.4c), which yields appτHyCbl = (1/ appkoff) = 200(120)s. Lastly, the time-traces of surface immobilized molecules clearly verify the previous
assumption that the KL kinetics, both in the presence and absence of ligand, are much faster thanthose associated with establishing a ligand binding equilibrium. In total, the collection of resultsfrom the above fluorescence biophysical studies provides unambiguous support for the proposedfour-state kinetic model (Figure 8.2) associated with the regulatory conformational transition of
the env8HyCbl riboswitch.
8.2.7

Relevance of kissing loop formation with respect to regulation of gene expression
To relate the biophysical behavior of this functional riboswitch with its biological

function, cell-based gene expression experiments (Figure 8.9) have been performed in an E.coli
strain lacking cobalamin adenosyl transferase (ΔbtuR), thereby preventing the subsequentconversion of hydroxocobalamin to adenosylcobalamin. In these experiments, cells weretransformed with vectors containing either the wild type or mutant riboswitches located upstreamof a GFPuv reporter gene—termed WT env8HyCbl+GFPuv or L5-mutant env8HyCbl+GFPuv,
respectively—and cultured in liquid media with or without 5000 nM HyCbl. The L5-mutant
env8HyCbl+GFPuv vector contains a riboswitch with the exact same dinucleotide substitution
used in the smFRET experiments (Supplemental Figure S8.1) and is predicted to be incapable ofregulating gene expression. In growth media lacking HyCbl, cells transformed with WT
env8HyCbl+GFPuv have normalized fluorescence values (see Experimental Methods) lower

216

than cells transformed with the L5-mutant env8HyCbl+GFPuv plasmid (grey bars, Figure 8.9b).
This nicely correlates with the biophysical observation that the WT env8HyCbl smFRET
construct is mostly undocked (FUfree ≈ 0.78) at 1 mM Mg2+, whereas the L5-mutant iscompletely undocked (FUfree ≈ 1.0). Additionally, when cultured in media containing 5000 nM
HyCbl, only cells harboring WT env8HyCbl+GFPuv show a substantial ligand-dependent
reduction in GFPuv expression (bronze bars, Figure 8.9b), which is well correlated with the
FUHyCbl = 0.3(1) observed in the smFRET experiments.

Figure 8.9 Cell-based regulation of gene expression via the env8HyCbl riboswitch. a) Schematic
diagram relating the four-state kinetic model to the regulation of gene expression in vivo. b)
Histograms comparing the normalized fluorescence and relative GFPuv expression of cell
cultures grown in the presence (bronze bars) or absence (grey bars) of 5000 nM HyCbl.

217

The high degree of correlation between the in vitro and in vivo experiments strongly suggests
that: (i) the fractional occupancy of the undocked conformation dictates the maximum level ofgene expression, regardless of whether or not ligand is bound and (ii) formation of the docked
conformation, resulting from a long-range KL interaction between L5 and L13, sequesters theribosome binding site within L13 and prevents initiation of translation.
8.3

Discussion
Ligand-induced conformational transitions are central to the function of riboswitches. A

significant body of structural work has revealed the numerous ways in which riboswitches can
specifically interact with their cognate ligands. However, much less is known about how
molecular recognition in the receptor domains translates into the regulatory domain’s ability toalter gene expression. The results of these single-molecule experiments mechanistically link: (i)conformational transitions facilitated by ligand binding to (ii) regulation of gene expression in a
fully functional riboswitch containing both of these domains. Specifically, the fluorescently
labeled RNA constructs examined in this work provide information about the equilibrium andkinetic behavior of a regulatory kissing loop (KL) interaction. Furthermore, the novel use ofhydroxocobalamin-induced fluorescence quenching as an orthogonal experimental observable
offers additional insights into the ligand binding process. Together, these results highlight several
key findings about the functional mechanism of gene expression for the env8HyCbl riboswitch.
The following is a discussion of these observations and their relevance to other areas of RNA
biology.
8.3.1

Kissing Loop docking dynamics
Both the freely diffusing and surface immobilized experiments clearly identify the L5-

L13 KL interaction as the structural motif that directly regulates biological function (i.e., gene218

expression). Additionally, this motif is also responsible for organizing the RNA’s global
conformation. Many other functional nucleic acids make use of similar types of tertiary
interactions to accomplish these same tasks. For example, KL are important for organizing thethree dimensional (3D) structure of self-cleaving ribozymes(26,27) and regulate control a varietyof biological processes(28), such as dimerization of retroviral genomes(22) and plasmid
replication(29,30). One characteristic of both intra- and inter-molecular KL interactions is theformation of canonical Watson-Crick base pairs between the two interacting loops. In addition to
5 complementary base pairs, the L5-L13 interaction contains a G-A mismatch adjacent to adinucleotide (AG) bulge. Presumably, these structural defects serve to destabilize thisinteraction. As a result, the KL will only be stably formed in the presence of HyCbl, thereby
conferring a ligand-dependent occlusion of the ribosome binding site. This clearly demonstrates
how nucleotide sequence can be used to tune the stability of a RNA kissing-loop. Other
examples exist where localization of Mg2+ modulates the stability of these types of interactions,
as is the case with intermolecular HIV (human immunodeficiency virus) genome dimerization(22). In the env8HyCbl riboswitch, divalent cations are essential for KL formation and likely
play a similar role. Indeed, the x-ray crystal structure of this RNA depicts substantial electron
density associated with Ba2+(the more strongly x-ray scattering mimetic of Mg2+) near theinteracting loops(18). While divalent cations and nucleotide sequence represent two effective
ways to adjust the stability of RNA KLs, there is also another ways to accomplish such a task.
8.3.2

KL docking in the presence of a macromolecular stabilizing agent
In addition to distinctly identifying the existence of a KL docking equilibrium in theabsence of ligand, the above experimental results reveal that KL docking and undocking also
occurs when HyCbl is bound. Specifically, the presence of a bound ligand alters the energetics of219

this process to substantially favor the docked conformation. This is yet another example of how
KL stability can be tuned for a particular biological function. Careful inspection of the HyCblbound crystal structure reveals that most of the intermolecular contacts are van der Waals
interactions governed by shape complementarity(18). These interactions significantly stabilize
the docked conformation, as evidenced by the ≈ 23-fold slower undocking rate constant when theligand is bound to the RNA. This is an excellent demonstration of how a specificmacromolecular cosolute can stabilize the formation of a KL interaction directly responsible fora specific biological function.
As a matter of fact, a similar mechanism is employed in the Rop-RNA I-RNA II plasmid
replication system(30). Specifically, the Rop protein acts as a molecular scaffold that stabilizes
the intermolecular KL interaction between the loops of RNA I and RNA II (31) and regulates
formation of RNA primers required for DNA plasmid replication. In this system, the presence of
Rop substantially reduces the rate at which RNA I dissociates from the ternary complex(32).
Interestingly, the six acetamide functional groups in the corrin ring of HyCbl chemicallyrecapitulate the amino acid side chains of glutamine and asparagine. One can speculate that theprotein-like chemical functionality of HyCbl allows this cofactor to imitate the stabilizing
properties of Rop. Moreover, Rop presents eight of these amino acid residues to the interface
between the protein and the kissing loops of RNA I and II(30,33). Certainly, the resounding
functional similarity between these two biological systems demonstrates how the formation of a
KL interaction can be facilitated by a macromolecular binding partner (e.g., protein or ligand) inorder to regulate a biologically relevant process (e.g., plasmid replication or gene expression).

2208.3.3

Ligand Recognition
A long-standing question in nucleic acid recognition of small molecule ligands iswhether: (i) an induced-fit or (ii) a conformational-capture model most appropriately describes
these aptamer-substrate interactions. The induced-fit model demands that ligand binding occurs
in open (e.g., undocked) conformations of the aptamer, which then induces a structural transition
that encapsulates the bound molecule(10). This is in contrast to the conformational-capture
model, whereby ligand recognition occurs in transient, highly organized folded structures,
similarly resulting in stabilization of the bound (e.g., docked) conformation(16). For theenv8HyCbl riboswitch, the experimental validation of the four-state model (Figure 8.6) and thesurface-immobilized time-traces in the presence of HyCbl (Figure 8.8a, Supplemental FigureS8.3) strongly suggest that ligand binding (i.e., quenching) occurs only in the undocked
conformation. This notion is further supported by existing structural data(18), which reveals thatthe env8HyCbl riboswitch conceals a majority (≈60%) of the solvent accessible surface area ofthe ligand when the KL interaction is formed. Therefore, the approximately spherical HyCbl
ligand would not be able to enter, or exit, the binding pocket when the L5-L13 interaction isformed. Additionally, isothermal titration calorimetry experiments demonstrate that riboswitcheslacking L13 can still effectively bind HyCbl, implying that ligand binding does readily occur
when the RNA is in the undocked conformation (18). Together, these observations support an
induced-fit ligand-recognition model for this riboswitch, which bolsters the notion that thisbehavior is commonly associated with many small-molecule binding RNAs(34-36).
Additionally, the ligand binding kinetics experiments reveal that the bimolecular appkbind
is fast (≥ 106 M-1s-1), relative to many other RNA-ligand interactions(37). However, therelatively slow (≤0.01 s-1) appkdiss mandates that, at HyCbl concentrations near appKd, the time-221scale associated with ligand binding (i.e., [appkon × appKd]-1 ≈ 200 s) is much longer than that
required to establish the two KL docking equilibria (i.e., [kdock,free + kundock,free]-1 ≈ 1 s and[kdock,HyCbl + kundock,HyCbl]-1 ≈ 0.1 s). Therefore, when the HyCbl concentration is less than ≈ 1 µM
(i.e., where there time-scale associated with these processes are similar, [appkbind × 1 µM]-1 ≈[kdock,free + kundock,free]-1), this uniquely identifies ligand binding as the rate limiting biophysicalprocess in the four-state kinetic model (Figure 8.2). This conclusion raises a significant question
with regard to genetic regulation: does the riboswitch-containing mRNA have enough time, after
being transcribed, to bind the ligand before being degraded in the cell? It is certainly worth
noting that, at HyCbl concentrations near appKd, the time-scale associated with ligand binding(i.e.,[appkon × appKd]-1 ≈ 200 s) is comparable to that of mRNA decay (≈ 300 s)(38). This would
seem to suggest that gene expression is somewhat temporally restricted. However, more rigorousand quantitative studies of mRNA translation and decay will be required to conclusively
determine the kinetic vs. thermodynamic nature of genetic regulation for the env8HyCbl
riboswitch(37).
8.3.4

Mechanism of regulation for the env8 hydroxocobalamin riboswitch
The following quantitative comparison of the in vitro and in vivo results provides a

substantial amount of information linking the physical mechanism of the env8HyCbl riboswitch
to the biologically relevant ligand-dependent modulation of gene expression. The L5-mutant
env8HyCbl+GFPuv transformed cells are shown to be insensitive to HyCbl (Figure 8.9b) and areused to define the maximum amount of gene expression resulting from completely undocked
RNAs. The normalized fluorescence (See Experimental Methods) associated with maximal gene
expression is used to calculate the relative gene expression associated with the WT env8HyCbl
riboswitch in the presence or absence of 5000 nM ligand. This quantitation of the cell-based222

experiments reveals that, in the absence of HyCbl, the env8HyCbl riboswitch encoded upstreamof a GFPuv fluorescence reporter results in a relative gene expression value of 0.90(8). This invivo value is nearly experimentally indistinguishable from the 1 mM Mg2+ in vitro values of
FUfree = 0.83(7) and FUfree ≈ 0.78 for surface-immobilized and freely diffusing smFRET
experiments, respectively. Such quantitative agreement indicates that the amount of gene
expressions is highly correlated with the FUfree. Furthermore, the WT env8HyCbl results in a
relative gene expression value of 0.10(3) in the presence of 5000 nM HyCbl. Again, this in vivo
value is similar to the 1 mM Mg2+ in vitro values of FUHyCbl = 0.3(1) and FUHyCbl ≈ 0.25
determined via surface-immobilized and freely diffusing smFRET experiments, respectively.
Together, the in vitro and in vivo experiments unambiguously demonstrate that the fractional
occupancy of the undocked conformation at 1mM Mg2+ is highly correlated with the relative
amount of GFPuv expression and that the L5-L13 KL interactions is the physical switch
associated with this regulatory element.
8.4

Conclusions
This work represents the first quantitative single-molecule investigations of a fully

functional riboswitch, complete with complementary cell-based assays. The results of theseexperiments clearly identify the conformational mechanism responsible for regulation of gene
expression by the env8HyCbl riboswitch. Specifically, ligand binding occurs when the L5-L13
kissing loop is absent and alters the energetics associated with this regulatory interaction,
resulting in preferential stabilization of the docked state. In this docked conformation, theribosome binding site (RBS) within L13 makes canonical Watson-Crick base pairing interactions
with L5. These base pairs conceal this region of the mRNA from the cellular machinery
responsible for translation initiation, thus repressing expression of the downstream gene. Time-223

resolved ligand-binding experiments indicate that although this process is fast relative to otherriboswitches, it is still much slower than the time-scale for equilibration of KL docking. The highdegree of correlation between cell-based GFPuv reporter assays and biophysical fluorescence
experiments further supports the notion that formation of the KL interaction is directly
responsible for repression of gene expression and identifies this region of the RNA as thephysical regulatory switch. This KL interaction shares a number of biochemical features withother functional nucleic acids. Together these similarities highlight the importance of nucleotide
sequence, divalent cations, and macromolecular binding partners in tuning the stability of thisstructural motif. One interesting direction would be to more rigorously explore the importance ofthe structural defects within the KL and their ability to confer ligand-facilitated docking. Lastly,and perhaps most importantly, this works presents a thorough and robust characterization of aligand-facilitated conformational transition that serves as a point-of-comparison for other suchstudies of similar RNAs.
8.5

Experimental Methods

8.5.1

Fluorescent Construct Design
All of the env8 riboswitch constructs used this in work are based on a sequence originallyidentified from comparative metagenomic analyses of functional non-coding RNAs(19). The
various constructs designed for single-molecule fluorescence resonance energy transfer(smFRET) experiments are prepared via non-splinted ligation(39) of two custom syntheticallymodified RNA oligonucleotides (Integrated DNA Technologies). The wild-type env8 smFRET
construct is assembled from the following two RNAs: WT strand 1; (5"-biotin-AAA AAA AAG
GCC UAA AAG CGU AGU GGG AAA G[dT*]G ACG UGA AAU UCG UCC AGA UUA C3") and WT strand 2; (5"-phosphate-UUG AUA CGG UUA UAC UCC GAA UGC CAC CUA

224

GGC CAU ACA ACG AGC AAG GAG ACU C-Cy3-3"). According to the x-ray crystal
structure of env8, nucleotide U24 is completely solvent exposed and lacks intramolecular
contacts with other functional groups of the RNA(18). By way of additional confirmation,
SHAPE chemical probing experiments reveal this nucleotide to be hyper-reactive, as is often thecase with highly solvent exposed nucleotides. In conjunction with the x-ray crystal structure, thisconclusion suggests that this position is well suited for synthetic modification, which will
negligibly impact the conformational dynamics. To this end, nucleotide U24 is replaced by anamino-modified nucleotide (dT*), which is subsequently chemically labeled with an NHSfunctionalized Cy5 fluorophore (GE healthcare) following the manufacturer’s suggested
protocol. Microcentrifuge size-exclusion columns (ThermoScientific) are used to remove theexcess unreacted fluorophores from the labeling reaction. The WT strands 1 and 2 are annealed
together by slowly cooling from 85 °C to room temperature in buffer (50 mM HEPES, 800 µM
hydroxocobalamin (Sigma Aldrich), 100 mM KCl, pH 7.5) prior to enzymatic ligation via T4
RNA ligase I (New England Biolabs). Isolation of full-length doubly labeled RNA construct isachieved through the use of a reverse phase high-performance liquid chromatography (HPLC)
column (Agilent).
Preparation of the L5-mutant construct is carried out using the same procedures expect that L5mutant strand 2 (5"-phosphate-AAG AUA CGG UUA UAC UCC GAA UGC CAC CUA GGC
CAU ACA ACG AGC AAG GAG ACU C-Cy3-3") is used instead of WT strand 2, with theunderlined nucleotides corresponding to the desired mutations. Synthesis of the Cy3-only env8
construct used in the ensemble fluorometry experiments is performed using in vitro transcription
by T7 RNA polymerase, as described previously(18). The resulting transcripts are exposed to
NaIO4 and NaBH3CN for selective oxidation of the 3"-ribose(40) and reacted with hydrazide-225

functionalized Cy3 to covalently attach the fluorophore to this position. Removal of excess dyeand HPLC purification is performed, as described above, in order to isolate full-length
fluorescently labeled RNA constructs.
8.5.2

Freely-Diffusing smFRET
Freely-diffusing smFRET experiments are performed on an home-build inverted confocal

fluorescence microscope, described previously(17), with a 1.2 N.A. water objective (Olympus
Corporation) in epifluorescence configuration to both focus the excitation light and collect theemitted fluorescence. When the concentration of fluorescently labeled RNA is sufficiently low(≈ 125 pM), individual molecules will transiently diffuse into the overlapping foci of two (532
nm,Time-Bandwidth products and 635 nm, PicoQuant GmbH) alternating laser excitation(41)(ALEX) sources with ≈ 100 µW average power. This yields a short (< 1 ms) burst offluorescence photons dictated by the time the RNA spends within the excitation and detection
volume. The FRET efficiency (EFRET) resulting from 532 nm excitation is calculated for eachfluorescent burst, where the use of ALEX techniques ensures that only dually labeled, nonphotophysically damaged constructs are considered for analysis. All experiments are performedat ambient temperatures (20-22 °C) in standard buffer (50 mM HEPES, 25 mM KOH, 10 mM
NaOH, 100 mM KCl, 2 mM TROLOX, 100 nM PCD and 5 mM PCA, pH 7.7) consisting of thewell-characterized enzymatic oxygen scavenging system, PCA/PCD(42), and various
concentrations of MgCl2 and hydroxocobalamin (HyCbl).
8.5.3

Ensemble Fluorometry
HyCbl-dependent Cy3 quenching ensemble experiments are performed on a fluorimeter(Jobin Yvon) with ≈ 500 pM fluorescent RNA in the standard buffer containing 1mM MgCl2 and10-50 nM hydroxocobalamin. The excitation and emission filters are set to 532 ± 1 and 570 ± 7226nm, respectively, with each data point representing one second of integrated fluorescence. The
narrow, off-peak excitation filter ensures that photobleaching of Cy3 represents a negligible
contribution to the time-dependent reduction in the fluorescence intensity.
8.5.4

Surface-Immobilized smFRET
Surface-immobilzed smFRET experiments are performed on the same inverted confocal

fluorescence microscope used for the freely-diffusing experiments. Immobilization isaccomplished using biotin-streptavidin chemistry, resulting in a surface coverage of < 1molecule per µm2. Average laser powers are ≈ 2 µW in order to reduce the effects ofphotobleaching and prolong the observation of surface immobilized molecules. Fluorescence
trajectories are constructed from 30 ms time bins of detected photons. Docking and undocking
rate constants are determined using either a previously described thresholding routine(43) or a
Bayesian maximum likelihood model(25). The two analyses result in rate constants that areexperimentally indistinguishable (when applied to the same data set).
8.5.5

in vivo Regulation of GFP Expression

Riboswitch constructs were amplified via recombinant overlapping PCR and cloned into a
pBR327 derivative between NsiI and HindIII restriction sites. For the cell-based assays, plasmids
are transformed into E.coli Keio strain ∆btuR (Keio collection(44) JW1262). The cells are plated
onto LB agar plates supplemented with 100 µg/mL ampicillin, and incubated at 37°C for 14-16
hours to facilitate colony formation. Three colonies are picked for each construct and grown tosaturation in a rich, chemically, defined medium (CSB media: NaH2PO4 (4.6 mg/mL), K2HPO4(11.7 mg/mL), and (NH4)2SO4 (2.0 mg/mL), D-glucose (0.6%), sodium citrate (5 mM), andMgSO4 (492 µM), FeCl3·6H2O (109 µM), ZnCl2·4H2O (28.3 µM), CoCl2·6H2O (24.8 µM),
Na2MoO4 (13.8 µM), CaCl2·2H2O (20.1 µM), CuCl2 (21.9 µM), MnCl2 (23.4 µM), and H3BO3

227

(23.9 µM)) supplemented with 100 µg/mL ampicillin to maintain plasmid selection. In order tomeasure cellular fluorescence resulting from GFPuv expression, 5 µL of saturated overnight
culture has been added to 5 mL of CSB medium supplemented with 100 µg/mL ampicillin and 5µM hydoxocobalamin, and grown to mid-log phase via incubation at 37°C in a rotating drum.
For fluorescence measurements, 300 µL of cells from each biological replicate are added to thewells of a Greiner 96 well, half-area microplate. GFPuv expression is monitored at an excitation
wavelength of 395 nm and a 510 nm emission wavelength using a Tecan Infinite M200 PRO
plate-reader. All data shown are the average fluorescence values of three biological replicatesmeasured in triplicate and normalized to cell density as determined by the OD600 for eachindividual well. Normalized fluorescence values for each construct are background corrected via
subtraction of cell-density normalized fluorescence from a pBR327 empty vector control.

228

8.6

References

1.

Tucker, B. J.; R. R. Breaker. Riboswitches as versatile gene control elements, Curr. Opin.
Struct. Biol., 2005, 15, 342-348.

2.

Nahvi, A.; N. Sudarsan; M. S. Ebert; X. Zou; K. L. Brown; R. R. Breaker. Genetic
control by a metabolite binding mRNA, Chem. Biol., 2002, 9, 1043.

3.

Nou, X.; R. J. Kadner. Adenosylcobalamin inhibits ribosome binding to btuB RNA,
Proc. Natl. Acad. Sci. USA, 2000, 97, 7190-7195.

4.

Banerjee, R.; S. W. Ragsdale. The many faces of vitamin B12: catalysis by cobalamindependent enzymes, Annu. Rev. Biochem., 2003, 72, 209-247.

5.

Nahvi, A.; J. E. Barrick; R. R. Breaker. Coenzyme B12 riboswitches are widespread
genetic control elements in prokaryotes, Nucleic Acids Res., 2004, 32, 143-150.

6.

Rodionov, D. A.; A. G. Vitreschak; A. A. Mironov; M. S. Gelfand. Comparative
genomics of the vitamin B12 metabolism and regulation in prokaryotes, J. Biol. Chem.,
2003, 278, 41148-41159.

7.

Martens, J. H.; H. Barg; M. J. Warren; D. Jahn. Microbial production of vitamin B12,
Appl. Microbiol. Biotechnol., 2002, 58, 275-285.

8.

Roth, J. R.; J. G. Lawrence; T. A. Bobik. Cobalamin (coenzyme B12): synthesis andbiological significance, Annu. Rev. Microbiol., 1996, 50, 137-181.

9.

Breaker, R. R. Riboswitches and the RNA world, Cold Spring Harb. Perspect. Biol.,2012, 4.

10.

Garst, A. D.; A. L. Edwards; R. T. Batey. Riboswitches: structures and mechanisms, Cold
Spring Harb. Perspect. Biol., 2011, 3.

11.

Roy, R.; S. Hohng; T. Ha. A Practical Guide to Single-Molecule FRET, Nat. Methods,
2008, 5, 507-516.

12.

Lemay, J. F.; J. C. Penedo; R. Tremblay; D. M. Lilley; D. A. Lafontaine. Folding of the
Adenine Riboswitch, Chem. Biol., 2006, 13, 857-868.

13.

Suddala, K. C.; A. J. Rinaldi; J. Feng; A. M. Mustoe; C. D. Eichhorn; J. A. Liberman; J.
E. Wedekind; H. M. Al-Hashimi; C. L. Brooks, 3rd; N. G. Walter. Single transcriptional
and translational preQ1 riboswitches adopt similar pre-folded ensembles that follow
distinct folding pathways into the same ligand-bound structure, Nucleic Acids Res., 2013,
41, 10462-10475.

14.

Souliere, M. F.; R. B. Altman; V. Schwarz; A. Haller; S. C. Blanchard; R. Micura.
Tuning a riboswitch response through structural extension of a pseudoknot, Proc. Natl.
Acad. Sci. USA, 2013, 110, E3256-3264.

229

15.

Haller, A.; R. B. Altman; M. F. Souliere; S. C. Blanchard; R. Micura. Folding and ligand
recognition of the TPP riboswitch aptamer at single-molecule resolution, Proc. Natl.
Acad. Sci. USA, 2013, 110, 4188-4193.

16.

Haller, A.; U. Rieder; M. Aigner; S. C. Blanchard; R. Micura. Conformational capture of
the SAM-II riboswitch, Nat. Chem. Biol., 2011, 7, 393-400.

17.

Fiegland, L. R.; A. D. Garst; R. T. Batey; D. J. Nesbitt. Single-molecule studies of thelysine riboswitch reveal effector-dependent conformational dynamics of the aptamer
domain, Biochemistry, 2012, 51, 9223-9233.

18.

Johnson, J. E., Jr.; F. E. Reyes; J. T. Polaski; R. T. Batey. B12 cofactors directly stabilize
an mRNA regulatory switch, Nature, 2012, 492, 133-137.

19.

Weinberg, Z.; J. X. Wang; J. Bogue; J. Yang; K. Corbino; R. H. Moy; R. R. Breaker.
Comparative genomics reveals 104 candidate structured RNAs from bacteria, archaea,
and their metagenomes, Genome Biol., 2010, 11, R31.

20.

Griffiths-Jones, S.; A. Bateman; M. Marshall; A. Khanna; S. R. Eddy. Rfam: an RNA
family database, Nucleic Acids Res., 2003, 31, 439-441.

21.

Xu, H.; Y. Li; C. Liu; Q. Wu; Y. Zhao; L. Lu; H. Tang. Fluorescence resonance energy
transfer between acridine orange and rhodamine 6G and its analytical application forvitamin B12 with flow-injection laser-induced fluorescence detection, Talanta, 2008, 77,
176-181.

22.

Ennifar, E.; P. Walter; B. Ehresmann; C. Ehresmann; P. Dumas. Crystal structures of
coaxially stacked kissing complexes of the HIV-1 RNA dimerization initiation site, Nat.
Struct. Biol., 2001, 8, 1064-1068.

23.

Horiya, S.; X. Li; G. Kawai; R. Saito; A. Katoh; K. Kobayashi; K. Harada. RNA LEGO:
magnesium-dependent formation of specific RNA assemblies through kissing
interactions, Chem. Biol., 2003, 10, 645-654.

24.

Tyrrell, J.; J. L. McGinnis; K. M. Weeks; G. J. Pielak. The cellular environment
stabilizes adenine riboswitch RNA structure, Biochemistry, 2013, 52, 8777-8785.

25.

Bronson, J. E.; J. Fei; J. M. Hofman; R. L. Gonzalez, Jr.; C. H. Wiggins. Learning rates
and states from biophysical time series: a Bayesian approach to model selection andsingle-molecule FRET data, Biophys. J., 2009, 97, 3196-3205.

26.

Andersen, A. A.; R. A. Collins. Intramolecular secondary structure rearrangement by the
kissing interaction of the Neurospora VS ribozyme, Proc. Natl. Acad. Sci. USA, 2001, 98,
7730-7735.

27.

Toor, N.; K. S. Keating; O. Fedorova; K. Rajashankar; J. Wang; A. M. Pyle. Tertiaryarchitecture of the Oceanobacillus iheyensis group II intron, RNA, 2010, 16, 57-69.

230

28.

Brunel, C.; R. Marquet; P. Romby; C. Ehresmann. RNA loop-loop interactions as
dynamic functional motifs, Biochimie, 2002, 84, 925-944.

29.

Wagner, E. G.; S. Brantl. Kissing and RNA stability in antisense control of plasmid
replication, Trends Biochem. Sci., 1998, 23, 451-454.

30.

Eguchi, Y.; J. Tomizawa. Complex formed by complementary RNA stem-loops and its
stabilization by a protein: function of CoIE1 Rom protein, Cell, 1990, 60, 199-209.

31.

Helmer-Citterich, M.; M. M. Anceschi; D. W. Banner; G. Cesareni. Control of ColE1
replication: low affinity specific binding of Rop (Rom) to RNAI and RNAII, Embo. J.,
1988, 7, 557-566.

32.

Di Primo, C. Real time analysis of the RNAI-RNAII-Rop complex by surface plasmon
resonance: from a decaying surface to a standard kinetic analysis, J. Mol. Recognit.,
2008, 21, 37-45.

33.

Banner, D. W.; M. Kokkinidis; D. Tsernoglou. Structure of the ColE1 rop protein at 1.7
A resolution, J. Mol. Biol., 1987, 196, 657-675.

34.

Williamson, J. R. Induced fit in RNA-protein recognition, Nat. Struct. Biol., 2000, 7,
834-837.

35.

Noeske, J.; J. Buck; B. Furtig; H. R. Nasiri; H. Schwalbe; J. Wohnert. Interplay of'induced fit' and preorganization in the ligand induced folding of the aptamer domain of
the guanine binding riboswitch, Nucleic Acids Res., 2007, 35, 572-583.

36.

Serganov, A.; Y. R. Yuan; O. Pikovskaya; A. Polonskaia; L. Malinina; A. T. Phan; C.
Hobartner; R. Micura; R. R. Breaker; D. J. Patel. Structural basis for discriminative
regulation of gene expression by adenine- and guanine-sensing mRNAs, Chem. Biol.,
2004, 11, 1729-1741.

37.

Zhang, J.; M. W. Lau; A. R. Ferre-D'Amare. Ribozymes and riboswitches: modulation of
RNA function by small molecules, Biochemistry, 2010, 49, 9123-9131.

38.

Bernstein, J. A.; A. B. Khodursky; P. H. Lin; S. Lin-Chao; S. N. Cohen. Global analysis
of mRNA decay and abundance in Escherichia coli at single-gene resolution using twocolor fluorescent DNA microarrays, Proc. Natl. Acad. Sci USA, 2002, 99, 9697-9702.

39.

Moore, M. J., Query, C.C. Uses of Site-Specifically Modified RNAs Constructed by
RNA Ligation, In RNA:Protein Interactions: A Practical Approach (Smith, C. W. J.,
Ed.), pp 75-140, Oxford University Press Inc., New York.1998

40.

Proudnikov, D.; A. Mirzabekov. Chemical methods of DNA and RNA fluorescentlabeling, Nucleic Acids Res., 1996, 24, 4535-4542.

41.

Kapanidis, A. N.; T. A. Laurence; N. K. Lee; E. Margeat; X. Kong; S. Weiss.
Alternating-laser excitation of single molecules, Acc. Chem. Res., 2005, 38, 523-533.

231

42.

Aitken, C. E.; R. A. Marshall; J. D. Puglisi. An Oxygen Scavenging System for
Improvement of Dye Stability in Single-Molecule Fluorescence Experiments, Biophys. J.,
2008, 94, 1826-1835.

43.

Hodak, J. H.; C. D. Downey; J. L. Fiore; A. Pardi; D. J. Nesbitt. Docking kinetics andequilibrium of a GAAA tetraloop-receptor motif probed by single-molecule FRET, Proc.
Natl. Acad. Sci. USA, 2005, 102, 10505–10510.

44.

Baba, T.; T. Ara; M. Hasegawa; Y. Takai; Y. Okumura; M. Baba; K. A. Datsenko; M.
Tomita; B. L. Wanner; H. Mori. Construction of Escherichia coli K-12 in-frame, singlegene knockout mutants: the Keio collection, Mol. Syst. Biol., 2006, 2, 2006 0008.

232

Bibliography
Chapter 1:
Introduction
1.

Dahm, R. Discovering DNA: Friedrich Miescher and the early years of nucleic acidresearch, Hum. Genet., 2008, 122, 565-581.

2.

Voet, D.; J. G. Voet (2011) Biochemistry, 4th ed., John Wiley & Sons, Hoboken, NJ.

3.

Crick, F. Central dogma of molecular biology, Nature, 1970, 227, 561-563.

4.

Silverman, S. K. (2006) Nucleic acid switches and sensors, Landes
Bioscience/Eurekah.com and Springer Science+Business Media, Georgetown, TX andNew York, NY.

5.

Watson, J. D.; F. H. Crick. Molecular structure of nucleic acids: a structure fordeoxyribose nucleic acid, Nature, 1953, 171, 737-738.

6.

Carvalho, J. C. Derivation of the mass of the observable universe, Int. J. Theor. Phys.,
1995, 34, 2507-2509.

7.

Holmstrom, E. D.; N. F. Dupuis; D. J. Nesbitt. Pulsed IR heating studies of singlemolecule DNA duplex dissociation kinetics and thermodynamics, Biophys. J., 2014, 106,220-231.

8.

Dupuis, N. F.; E. D. Holmstrom; D. J. Nesbitt. Single-Molecule Kinetics Reveal CationPromoted DNA Duplex Formation Through Ordering of Single-Stranded Helices,
Biophys. J., 2013, 105, 756-766.

9.

Hodak, J. H.; C. D. Downey; J. L. Fiore; A. Pardi; D. J. Nesbitt. Docking kinetics andequilibrium of a GAAA tetraloop-receptor motif probed by single-molecule FRET, Proc.
Natl. Acad. Sci. USA, 2005, 102, 10505–10510.

10.

Holmstrom, E. D.; D. J. Nesbitt. Single Molecule FRET Studies of the Human
Telomerase RNA Pseudoknot: Temperature/Urea Dependent Folding Kinetics andThermodynamics, J. Phys. Chem. B., 2014.

11.

Fiore, J. L.; E. D. Holmstrom; L. R. Fiegland; J. H. Hodak; D. J. Nesbitt. The Role ofCounterion Valence and Size in GAAA Tetraloop-Receptor Docking/Undocking
Kinetics, J. Mol. Biol., 2012.

12.

Draper, D. E.; D. Grilley; A. M. Soto. Ions and RNA folding, Annu. Rev. Biophys.
Biomolec. Struct., 2005, 34, 221–243.

13.

Leipply, D.; D. Lambert; D. E. Draper. Ion-RNA interactions: Thermodynamic analysisof the effects of mono- and divalent ions on RNA conformational equilibria, Methods
Enzymol., 2009, 469, 433-463.

233

14.

Holmstrom, E. D.; J. L. Fiore; D. J. Nesbitt. Thermodynamic origins of monovalentfacilitated RNA folding, Biochemistry, 2012, 51, 3732-3743.

15.

Shelton, V. M.; T. R. Sosnick; T. Pan. Applicability of Urea in the Thermodynamic
Analysis of Secondary and Tertiary RNA Folding, Biochemistry, 1999, 38, 16831-16839.

16.

Lambert, D.; D. E. Draper. Effects of Osmolytes on RNA Secondary and Tertiary
Structure Stabilities and RNA-Mg2+ Interactions, J. Mol. Biol., 2007, 370, 993-1005.

17.

Lambert, D.; D. E. Draper. Denaturation of RNA secondary and tertiary structure byurea: simple unfolded state models and free energy parameters account for measured mvalues, Biochemistry, 2012, 51, 9014-9026.

18.

Priyakumar, U. D.; C. Hyeon; D. Thirumalai; A. D. MacKerell. Urea Destabilizes RNA
by Forming Stacking Interactions and Multiple Hydrogen Bonds with Nucleic Acid
Bases, J. Am. Chem. Soc., 2009, 131, 17759-17761.

19.

Lambert, D.; D. Leipply; D. E. Draper. The osmolyte TMAO stabilizes native RNA
tertiary structures in the absence of Mg2+: Evidence for a large barrier to folding from
phosphate dehydration, J. Mol. Biol., 2010, 404, 138-157.

20.

Breaker, R. R. Riboswitches and the RNA world, Cold Spring Harb. Perspect. Biol.,2012, 4.

21.

Garst, A. D.; A. L. Edwards; R. T. Batey. Riboswitches: structures and mechanisms, Cold
Spring Harb. Perspect. Biol., 2011, 3.

22.

Kador, L.; D. E. Horne; W. E. Moerner. Optical-Detection and Probing of Single Dopant
Molecules of Pentacene in a P-Terphenyl Host Crystal by Means of AbsorptionSpectroscopy, J. Phys. Chem., 1990, 94, 1237-1248.

23.

Moerner, W. E.; L. Kador. Optical-Detection and Spectroscopy of Single Molecules in a
Solid, Phys. Rev. Lett., 1989, 62, 2535-2538.

24.

Orrit, M.; J. Bernard. Single pentacene molecules detected by fluorescence excitation in ap-terphenyl crystal, Phys. Rev. Lett., 1990, 65, 2716-2719.

25.

Joo, C.; H. Balci; Y. Ishitsuka; C. Buranachai; T. Ha. Advances in single-molecule
fluorescence methods for molecular biology, Annu. Rev. Biochem., 2008, 77, 51-76.

26.

Roy, R.; S. Hohng; T. Ha. A Practical Guide to Single-Molecule FRET, Nat. Methods,
2008, 5, 507-516.

27.

Zhao, R.; D. Rueda. RNA folding dynamics by single-molecule fluorescence resonanceenergy transfer, Methods, 2009, 49, 112-117.

28.

Förster, T. Zwischenmolekulare Energiewanderung und Fluoreszenz, 1948, 437, 55-75.

234

29.

Einstein, A. Strahlungs-Emission und -Absorption nach der Quantentheorie, Deutch.
Phys. Ges., 1916, 18, 318-323.

30.

Schuler, B.; E. A. Lipman; P. J. Steinbach; M. Kumke; W. A. Eaton. Polyproline and the"spectroscopic ruler" revisited with single-molecule fluorescence, Proc. Natl. Acad. Sci.
USA, 2005, 102, 2754-2759.

31.

Akiyama, B. M.; M. D. Stone. Assembly of Complex RNAs by Splinted Ligation,
Methods Enzymol., 2009, 469, 27-46.

32.

Fiore, J. L.; E. D. Holmstrom; D. J. Nesbitt. Entropic Origin of Mg2+-Facilitated RNA
Folding, Proc. Natl. Acad. Sci. USA, 2012, 109, 2902-2907.

33.

Fiore, J. L.; B. Kraemer; F. Koberling; R. Erdmann; D. J. Nesbitt. Enthalpy-Driven RNA
Folding: Single-Molecule Thermodynamics of Tetraloop–Receptor Tertiary Interaction,
Biochemistry, 2009, 48, 2550–2558.

34.

Holmstrom, E. D.; D. J. Nesbitt. Real-time infrared overtone laser control of temperaturein picoliter H2O samples: "Nanobathtubs" for single-molecule microscopy, J. Phys.
Chem. Lett., 2010, 1, 2264–2268.
Chapter 2:
Experimental Methods

1.

Einstein, A. Strahlungs-Emission und -Absorption nach der Quantentheorie, Deutch.
Phys. Ges., 1916, 18, 318-323.

2.

van der Meer, B. W.; D. M. van der Meer; S. S. Vogel. Optimizing the Orientation Factor
Kappa-Squared for More Accurate FRET Measurements, In FRET – Förster Resonance
Energy Transfer, pp 63-104, Wiley-VCH Verlag GmbH & Co. KGaA.2013

3.

Fiore, J. L. (2012) Single-Molecule Kinetic and Thermodynamic Studies of CationFacilitated RNA Folding: Isolated Tertiary Interactions, In Chemical Physics, Universityof Colorado, Boulder.

4.

Baker, T. A. (2012) Confocal Microscopy Studies of Fluorescence Blinking of
Semiconductor Quantum Dots, Metal Nanoparticle Photogeneration, and Multiphoton
Photoemission from Thin Metal Films and Metal Nanoparticles, In Chemistry, Universityof Colorado, Boulder.

5.

Kapanidis, A. N.; T. A. Laurence; N. K. Lee; E. Margeat; X. Kong; S. Weiss.
Alternating-laser excitation of single molecules, Acc. Chem. Res., 2005, 38, 523-533.

6.

Kapanidis, A. N.; N. K. Lee; T. A. Laurence; S. Doose; E. Margeat; S. Weiss.
Fluorescence-aided molecule sorting: Analysis of structure and interactions byalternating-laser excitation of single molecules, Proc. Natl. Acad. Sci. USA, 2004, 101,
8936-8941.
235

7.

Lee, N. K.; A. N. Kapanidis; H. R. Koh; Y. Korlann; S. O. Ho; Y. Kim; N. Gassman; S.
K. Kim; S. Weiss. Three-color alternating-laser excitation of single molecules:
Monitoring multiple interactions and distances, Biophys. J., 2007, 92, 303–312.

8.

Lee, N. K.; A. N. Kapanidis; Y. Wang; X. Michalet; J. Mukhopadhyay; R. H. Ebright; S.
Weiss. Accurate FRET Measurements within Single Diffusing Biomolecules using
Alternating-Laser Excitation, Biophys. J., 2005, 88, 2939-2953.

9.

Forconi, M.; D. Herschlag. Metal ion-based RNA cleavage as a structural probe, Methods
Enzymol., 2009, 468, 91-106.

10.

Lonnberg, T.; M. Luomala. Intracomplex general acid/base catalyzed cleavage of RNA
phosphodiester bonds: the leaving group effect, Org. Biomol. Chem., 2012, 10, 67856791.

11.

Rasnik, I.; S. A. McKinney; T. Ha. Nonblinking and long-lasting single-molecule
fluorescence imaging, Nat. Methods, 2006, 3, 891-893.

12.

Aitken, C. E.; R. A. Marshall; J. D. Puglisi. An Oxygen Scavenging System for
Improvement of Dye Stability in Single-Molecule Fluorescence Experiments, Biophys. J.,
2008, 94, 1826-1835.

13.

Ruttinger, S.; V. Buschmann; B. Kramer; R. Erdmann; R. Macdonald; F. Koberling.
Comparison and accuracy of methods to determine the confocal volume for quantitative
fluorescence correlation spectroscopy, J. Microsc. Oxford, 2008, 232, 343-352.

14.

Joo, C.; H. Balci; Y. Ishitsuka; C. Buranachai; T. Ha. Advances in single-molecule
fluorescence methods for molecular biology, Annu. Rev. Biochem., 2008, 77, 51-76.

15.

Torella, J. P.; S. J. Holden; Y. Santoso; J. Hohlbein; A. N. Kapanidis. Identifying
molecular dynamics in single-molecule FRET experiments with burst variance analysis,
Biophys. J., 2011, 100, 1568-1577.

16.

Fiore, J. L.; J. H. Hodak; O. Piestert; C. D. Downey; D. J. Nesbitt. Monovalent andDivalent Promoted GAAA-Tetraloop–Receptor Tertiary Interactions from Freely
Diffusing Single-Molecule Studies, Biophys. J., 2008, 95, 3892-3905.

Chapter 3:
Real-Time IR Overtone Laser Control of Temperature in Picoliter H2O Samples: “NanoBathtubs” for Single Molecule Microscopy”
1.

Li, S.; K. Zhang; J. M. Yang; L. W. Lin; H. Yang. Single quantum dots as local
temperature markers, Nano Lett, 2007, 7, 3102–3105.

2.

Valerini, D.; A. Creti; M. Lomascolo; L. Manna; R. Cingolani; M. Anni. Temperaturedependence of the photoluminescence properties of colloidal CdSe/ZnS core/shell
quantum dots embedded in a polystyrene matrix, Phys. Rev. B, 2005, 71, 235409.
236

3.

Bartley, L. E.; X. W. Zhuang; R. Das; S. Chu; D. Herschlag. Exploration of theTransition State for Tertiary Structure Formation Between an RNA Helix and a Large
Structured RNA, J. Mol. Biol., 2003, 328, 1011–1026.

4.

Hohng, S.; T. J. Wilson; E. Tan; R. M. Clegg; D. M. J. Lilley; T. J. Ha. Conformational
flexibility of four-way junctions in RNA, J. Mol. Biol., 2004, 336, 69–79.

5.

Kato, H.; T. Nishizaka; T. Iga; K. Kinosita, Jr.; S. Ishiwata. Imaging of thermal activationof actomyosin motors, Proc. Natl. Acad. Sci. USA, 1999, 96, 9602–9606.

6.

de Mello, A. J.; M. Habgood; N. L. Lancaster; T. Welton; R. C. R. Wootton. Precise
temperature control in microfluidic devices using Joule heating of ionic liquids, Lab on a
Chip, 2004, 4, 417-419.

7.

Hu, G. Q.; Q. Xiang; R. Fu; B. Xu; R. Venditti; D. Q. Li. Electrokinetically controlled
real-time polymerase chain reaction in microchannel using Joule heating effect, Anal.
Chim. Acta, 2006, 557, 146–151.

8.

Zhao, Q. L.; Q. A. Huang; Y. C. Lin. Design and numerical analysis of a joule-heatinginduced continuous-flow polymerase chain reaction microchip, J. Micro-Nanolithogr.
MEMs and MOEMs, 2009, 8, 021101.

9.

Braun, D.; A. Libchaber. Trapping of DNA by thermophoretic depletion and convection,
Phys. Rev. Lett., 2002, 89, 188103.

10.

Ebbinghaus, S.; A. Dhar; D. McDonald; M. Gruebele. Protein folding stability anddynamics imaged in a living cell, Nat. Methods, 2010, 7, 319-323.

11.

Schoen, I.; H. Krammer; D. Braun. Hybridization kinetics is different inside cells, Proc.
Natl. Acad. Sci. USA., 2009, 106, 21649-21654.

12.

Zhao, R.; E. Alemán; D. Rueda. Laser Assisted Single-molecule Refolding, Biophys. J.,
2009, 96, Biophys. Soc. proceedings 9a.

13.

Slyadnev, M. N.; Y. Tanaka; M. Tokeshi; T. Kitamori. Photothermal temperature control
of a chemical reaction on a microchip using an infrared diode laser, Anal. Chem., 2001,
73, 4037–4044.

14.

Mao, H. B.; J. R. Arias-Gonzalez; S. B. Smith; I. Tinoco; C. Bustamante. Temperature
control methods in a laser tweezers system, Biophys. J., 2005, 89, 1308–1316.

15.

Peterman, E. J. G.; F. Gittes; C. F. Schmidt. Laser-induced heating in optical traps,
Biophys. J., 2003, 84, 1308–1316.

16.

Fiore, J. L.; B. Kraemer; F. Koberling; R. Erdmann; D. J. Nesbitt. Enthalpy-Driven RNA
Folding: Single-Molecule Thermodynamics of Tetraloop–Receptor Tertiary Interaction,
Biochemistry, 2009, 48, 2550–2558.

237

17.

Wieliczka, D. M.; S. S. Weng; M. R. Querry. Wedge shaped cell for highly absorbent
liquids: Infrared optical constants of water, Appl. Opt., 1989, 28, 1714–1719.

18.

Karstens, T.; K. Kobs. Rhodamine B and rhodamine 101 as reference substances forfluorescence quantum yield measurements, J. Phys. Chem., 1980, 84, 1871–1872.

19.

Muller, C. B.; K. Weiss; A. Loman; J. Enderlein; W. Richtering. Remote temperature
measurements in femto-liter volumes using dual-focus-Fluorescence Correlation
Spectroscopy, Lab on a Chip, 2009, 9, 1248-1253.

20.

Robinson, T.; Y. Schaerli; R. Wootton; F. Hollfelder; C. Dunsby; G. Baldwin; M. Neil;
P. French; A. Demello. Removal of background signals from fluorescence thermometry
measurements in PDMS microchannels using fluorescence lifetime imaging, Lab on a
Chip, 2009, 9, 3437–3441.

21.

Ross, D.; M. Gaitan; L. E. Locascio. Temperature measurement in microfluidic systems
using a temperature-dependent fluorescent dye, Anal. Chem., 2001, 73, 4117–4123.

22.

Arbeloa, F. L.; T. L. Arbeloa; M. J. T. Estevez; I. L. Arbeloa. Photophysics of
Rhodamines - Molecular-Structure and Solvent Effects, J. Phys. Chem., 1991, 95, 2203–
2208.

23.

Casey, K. G.; E. L. Quitevis. Effect of solvent polarity on nonradiative processes inxanthene dyes: Rhodamine B in normal alcohols, J. Phys. Chem., 1988, 92, 6590.

24.

Downey, C. D.; J. L. Fiore; C. D. Stoddard; J. H. Hodak; D. J. Nesbitt; A. Pardi. Metal
ion dependence, thermodynamics, and kinetics for intramolecular docking of a GAAA
tetraloop and receptor connected by a flexible linker, Biochemistry, 2006, 45, 3664–3673.

Chapter 4:
Pulsed Infrared-Heating Studies of Single Molecule DNA Duplex Dissociation Kinetics andThermodynamics
1.

Joo, C.; H. Balci; Y. Ishitsuka; C. Buranachai; T. Ha. Advances in single-molecule
fluorescence methods for molecular biology, Annu. Rev. Biochem., 2008, 77, 51-76.

2.

Roy, R.; S. Hohng; T. Ha. A Practical Guide to Single-Molecule FRET, Nat. Methods,
2008, 5, 507-516.

3.

Zhao, R.; D. Rueda. RNA folding dynamics by single-molecule fluorescence resonanceenergy transfer, Methods, 2009, 49, 112-117.

4.

Holmstrom, E. D.; J. L. Fiore; D. J. Nesbitt. Thermodynamic origins of monovalentfacilitated RNA folding, Biochemistry, 2012, 51, 3732-3743.

238

5.

Tarsa, P. B.; R. R. Brau; M. Barch; J. M. Ferrer; Y. Freyzon; P. Matsudaira; M. J. Lang.
Detecting force-induced molecular transitions with fluorescence resonant energy transfer,
Angew. Chem. Int. Ed., 2007, 46, 1999-2001.

6.

Shroff, H.; B. M. Reinhard; M. Siu; H. Agarwal; A. Spakowitz; J. Liphardt.
Biocompatible force sensor with optical readout and dimensions of 6 nm(3), Nano Lett.,
2005, 5, 1509-1514.

7.

Fiore, J. L.; B. Kraemer; F. Koberling; R. Erdmann; D. J. Nesbitt. Enthalpy-Driven RNA
Folding: Single-Molecule Thermodynamics of Tetraloop–Receptor Tertiary Interaction,
Biochemistry, 2009, 48, 2550–2558.

8.

Graneli, A.; C. C. Yeykal; R. B. Robertson; E. C. Greene. Long-distance lateral diffusionof human Rad51 on double-stranded DNA, Proc. Natl. Acad. Sci. USA, 2006, 103, 12211226.

9.

Fiore, J. L.; E. D. Holmstrom; D. J. Nesbitt. Entropic Origin of Mg2+-Facilitated RNA
Folding, Proc. Natl. Acad. Sci. USA, 2012, 109, 2902-2907.

10.

Zhao, R.; M. Marshall; E. A. Aleman; R. Lamichhane; A. Feig; D. Rueda. Laser-assisted
single-molecule refolding (LASR), Biophys. J., 2010, 99, 1925-1931.

11.

Gross, P.; G. Farge; E. J. G. Peterman; G. J. L. Wuite. Combining optical tweezers,
single-molecule fluorescence microscopy, and microfluidics for studies of DNA-protein
interactions, Method Enzymol., 2010, 475, 427-453.

12.

Holmstrom, E. D.; D. J. Nesbitt. Real-time infrared overtone laser control of temperaturein picoliter H2O samples: "Nanobathtubs" for single-molecule microscopy, J. Phys.
Chem. Lett., 2010, 1, 2264–2268.

13.

Braun, D.; A. Libchaber. Trapping of DNA by thermophoretic depletion and convection,
Phys. Rev. Lett., 2002, 89, 188103.

14.

Slyadnev, M. N.; Y. Tanaka; M. Tokeshi; T. Kitamori. Photothermal temperature control
of a chemical reaction on a microchip using an infrared diode laser, Anal. Chem., 2001,
73, 4037–4044.

15.

Mao, H. B.; J. R. Arias-Gonzalez; S. B. Smith; I. Tinoco; C. Bustamante. Temperature
control methods in a laser tweezers system, Biophys. J., 2005, 89, 1308–1316.

16.

Peterman, E. J. G.; F. Gittes; C. F. Schmidt. Laser-induced heating in optical traps,
Biophys. J., 2003, 84, 1308–1316.

17.

Zondervan, R.; F. Kulzer; H. van der Meer; J. A. Disselhorst; M. Orrit. Laser-driven
microsecond temperature cycles analyzed by fluorescence polarization microscopy,
Biophys. J., 2006, 90, 2958-2969.

239

18.

Yuan, H.; T. Xia; B. Schuler; M. Orrit. Temperature-cycle single-molecule FRET
microscopy on polyprolines, Phys. Chem. Chem. Phys., 2011, 13, 1762-1769.

19.

Wieliczka, D. M.; S. S. Weng; M. R. Querry. Wedge shaped cell for highly absorbent
liquids: Infrared optical constants of water, Appl. Opt., 1989, 28, 1714–1719.

20.

Hung, M. S.; O. Kurosawa; M. Washizu. Single DNA molecule denaturation using laserinduced heating, Mol. Cell. Probes, 2012, 26, 107-112.

21.

Holmberg, A.; A. Blomstergren; O. Nord; M. Lukacs; J. Lundeberg; M. Uhlen. The
biotin-streptavidin interaction can be reversibly broken using water at elevatedtemperatures, Electrophoresis, 2005, 26, 501-510.

22.

Sanborn, M. E.; B. K. Connolly; K. Gurunathan; M. Levitus. Fluorescence properties andphotophysics of the sulfoindocyanine Cy3 linked covalently to DNA, J. Phys. Chem. B.,
2007, 111, 11064-11074.

23.

Marras, S. A. Selection of fluorophore and quencher pairs for fluorescent nucleic acidhybridization probes, Methods Mol. Biol., 2006, 335, 3-16.

24.

Hyre, D. E.; I. Le Trong; E. A. Merritt; J. F. Eccleston; N. M. Green; R. E. Stenkamp; P.
S. Stayton. Cooperative hydrogen bond interactions in the streptavidin-biotin system,
Protein Sci., 2006, 15, 459-467.

25.

Selvin, P. R.; T. Ha, (Eds.) (2008) Single-molecule Techniques: A Laboratory Manual,
Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.

26.

Lamichhane, R.; A. Solem; W. Black; D. Rueda. Single-molecule FRET of proteinnucleic acid and protein-protein complexes: Surface passivation and immobilization,
Methods, 2010, 52, 192-200.

27.

Akabayov, B.; A. Henn; G. Nautrup-Pedersen; M. Elbaum; I. Sagi. (2002)
Immobilization of RNA: application to single molecule spectroscopy, In Molecular,
Cellular and Tissue Engineering, 2002. Proc. IEEE-EMBS Special Topic Conference, pp71-72.

28.

Janissen, R.; L. Oberbarnscheidt; F. Oesterhelt. Optimized straight forward procedure forcovalent surface immobilization of different biomolecules for single molecule
applications, Colloid. Surface B., 2009, 71, 200-207.

29.

Aleman, E. A.; H. S. Pedini; D. Rueda. Covalent-bond-based immobilization approaches
for single-molecule fluorescence, ChemBioChem, 2009, 10, 2862-2866.

30.

Schwartz, J. J.; S. Stavrakis; S. R. Quake. Colloidal lenses allow high-temperature singlemolecule imaging and improve fluorophore photostability, Nat. Nanotechnol., 2010, 5,
127-132.

240

31.

Altman, R. B.; D. S. Terry; Z. Zhou; Q. Zheng; P. Geggier; R. A. Kolster; Y. Zhao; J. A.
Javitch; J. D. Warren; S. C. Blanchard. Cyanine fluorophore derivatives with enhanced
photostability, Nat. Methods, 2012, 9, 68-71.

32.

Chen, Y. Y.; A. W. Wood. Application of a temperature-dependent fluorescent dye(Rhodamine B) to the measurement of radiofrequency radiation-induced temperaturechanges in biological samples, Bioelectromagnetics, 2009, 30, 583-590.

33.

Karstens, T.; K. Kobs. Rhodamine B and rhodamine 101 as reference substances forfluorescence quantum yield measurements, J. Phys. Chem., 1980, 84, 1871–1872.

34.

Casey, K. G.; E. L. Quitevis. Effect of solvent polarity on nonradiative processes inxanthene dyes: Rhodamine B in normal alcohols, J. Phys. Chem., 1988, 92, 6590.

35.

Maus, M.; M. Cotlet; J. Hofkens; T. Gensch; F. C. De Schryver; J. Schaffer; C. A. M.
Seidel. An experimental comparison of the maximum likelihood estimation and nonlinear
least squares fluorescence lifetime analysis of single molecules, Anal. Chem., 2001, 73,
2078-2086.

36.

Nesbitt, D. J.; N. F. Dupuis; E. D. Holstrom. Single Molecule Studies of StreptavidinBiotin Dissociation at Elevated Temperatures, Biophys. J., 2012, 102, 270a.

37.

Lucius, A. L.; N. K. Maluf; C. J. Fischer; T. M. Lohman. General methods for analysis ofsequential "n-step" kinetic mechanisms: application to single turnover kinetics ofhelicase-catalyzed DNA unwinding, Biophys. J., 2003, 85, 2224-2239.

38.

Park, J.; S. Myong; A. Niedziela-Majka; K. S. Lee; J. Yu; T. M. Lohman; T. Ha. PCRA
helicase dismantles RecA filaments by reeling in DNA in uniform steps, Cell, 2010, 142,544-555.

39.

Ragunathan, K.; C. Joo; T. Ha. Real-time observation of strand exchange reaction withhigh spatiotemporal resolution, Structure, 2011, 19, 1064-1073.

40.

Watson, J. D.; F. H. C. Crick. Molecular structure of nucleic acids - a structure fordeoxyribose nucleic acid, Nature, 1953, 171, 737-738.

41.

Porschke, D.; O. C. Uhlenbeck; F. H. Martin. Thermodynamics and kinetics of helix-coiltransition of oligomers containing GC base pairs, Biopolymers, 1973, 12, 1313-1335.

42.

Dewey, T. G.; D. H. Turner. Laser temperature jump study of solvent effects onpoly(adenylic acid) stacking, Biochemistry, 1980, 19, 1681-1685.

43.

Freier, S. M.; K. O. Hill; T. G. Dewey; L. A. Marky; K. J. Breslauer; D. H. Turner.
Solvent Effects on the Kinetics and Thermodynamics of Stacking in Poly(Cytidylic
Acid), Biochemistry, 1981, 20, 1419-1426.

44.

Dewey, T. G.; D. H. Turner. Laser temperature-jump study of stacking in adenylic acidpolymers, Biochemistry, 1979, 18, 5757-5762.

241

45.

Ikuta, S.; K. Takagi; R. B. Wallace; K. Itakura. Dissociation kinetics of 19 base paired
oligonucleotide-DNA duplexes containing different single mismatched base pairs,
Nucleic Acids Res., 1987, 15, 797-811.

46.

Sauer-Budge, A. F.; J. A. Nyamwanda; D. K. Lubensky; D. Branton. Unzipping kineticsof double-stranded DNA in a nanopore, Phys. Rev. Lett., 2003, 90, 238101.

47.

Liphardt, J.; B. Onoa; S. B. Smith; I. Tinoco, Jr.; C. Bustamante. Reversible unfolding of
single RNA molecules by mechanical force, Science, 2001, 292, 733-737.

48.

Biancaniello, P. L.; A. J. Kim; J. C. Crocker. Long-time stretched exponential kinetics insingle DNA duplex dissociation, Biophys. J., 2008, 94, 891-896.

49.

Gunnarsson, A.; P. Jonsson; V. P. Zhdanov; F. Hook. Kinetic and thermodynamic
characterization of single-mismatch discrimination using single-molecule imaging,
Nucleic Acids Res., 2009, 37, e99.

50.

Tawa, K.; W. Knoll. Mismatching base-pair dependence of the kinetics of DNA-DNA
hybridization studied by surface plasmon fluorescence spectroscopy, Nucleic Acids Res.,
2004, 32, 2372-2377.

51.

Sinden, R. R. (1994) DNA Structure and Function, Academic Press, San Diego, CA.

52.

Yakovchuk, P.; E. Protozanova; M. D. Frank-Kamenetskii. Base-stacking and basepairing contributions into thermal stability of the DNA double helix, Nucleic Acids Res.,
2006, 34, 564-574.

53.

Porschke, D. A direct measurement of the unzippering rate of a nucleic acid double helix,
Biophys. Chem., 1974, 2, 97-101.

54.

Nikolova, E. N.; H. M. Al-Hashimi. Preparation, resonance assignment, and preliminary
dynamics characterization of residue specific 13C/15N-labeled elongated DNA for the
study of sequence-directed dynamics by NMR, J. Biomol. NMR, 2009, 45, 9-16.

55.

Porschke, D. Model calculations on the kinetics of oligonucleotide double helix coiltransitions. Evidence for a fast chain sliding reaction, Biophys. Chem., 1974, 2, 83-96.

56.

Dupuis, N. F.; E. D. Holmstrom; D. J. Nesbitt. Single-Molecule Kinetics Reveal CationPromoted DNA Duplex Formation Through Ordering of Single-Stranded Helices,
Biophys. J., 2013, 105, 756-766.

57.

SantaLucia, J. A unified view of polymer, dumbbell, and oligonucleotide DNA nearestneighbor thermodynamics, Proc. Natl. Acad. Sci. USA, 1998, 95, 1460-1465.

58.

Hodak, J. H.; C. D. Downey; J. L. Fiore; A. Pardi; D. J. Nesbitt. Docking kinetics andequilibrium of a GAAA tetraloop-receptor motif probed by single-molecule FRET, Proc.
Natl. Acad. Sci. USA, 2005, 102, 10505–10510.

242

59.

Aitken, C. E.; R. A. Marshall; J. D. Puglisi. An Oxygen Scavenging System for
Improvement of Dye Stability in Single-Molecule Fluorescence Experiments, Biophys. J.,
2008, 94, 1826-1835.

60.

Kapanidis, A. N.; T. A. Laurence; N. K. Lee; E. Margeat; X. Kong; S. Weiss.
Alternating-laser excitation of single molecules, Acc. Chem. Res., 2005, 38, 523-533.

61.

Lee, N. K.; A. N. Kapanidis; Y. Wang; X. Michalet; J. Mukhopadhyay; R. H. Ebright; S.
Weiss. Accurate FRET Measurements within Single Diffusing Biomolecules using
Alternating-Laser Excitation, Biophys. J., 2005, 88, 2939-2953.
Chapter 5:
Thermodynamic Origins of Monovalent-Facilitated RNA Folding

1.

Mattick, J. S.; I. V. Makunin. Non-coding RNA, Hum. Mol. Genet., 2006, 15, R17-R29.

2.

Butcher, S. E.; A. M. Pyle. The molecular interactions that stabilize RNA tertiary
structure: RNA motifs, patterns, and networks, Acc. Chem. Res., 2011, 44, 1302-1311.

3.

Ditzler, M. A.; D. Rueda; J. J. Mo; K. Hakansson; N. G. Walter. A rugged free energy
landscape separates multiple functional RNA folds throughout denaturation, Nucl. Acids
Res., 2008, 36, 7088-7099.

4.

Steiner, M.; D. Rueda; R. K. O. Sigel. Ca2+ induces the formation of two distinct
subpopulations of group II intron molecules, Angew. Chem. Int. Ed. Engl. , 2009, 48,
9739-9742.

5.

Xie, Z.; N. Srividya; T. R. Sosnick; T. Pan; N. F. Scherer. Single-molecule studies
highlight conformational heterogeneity in the early folding steps of a large ribozyme,
Proc. Natl. Acad. Sci. USA, 2004, 101, 534-539.

6.

Zhuang, X. W.; H. Kim; M. J. B. Pereira; H. P. Babcock; N. G. Walter; S. Chu.
Correlating structural dynamics and function in single ribozyme molecules, Science,
2002, 296, 1473–1476.

7.

Record, M. T.; W. T. Zhang; C. F. Anderson. Analysis of effects of salts and uncharged
solutes on protein and nucleic acid equilibria and processes: A practical guide torecognizing and interpreting polyelectrolyte effects, Hofmeister effects, and osmotic
effects of salts, Adv. Protein. Chem., 1998, 51, 281-353.

8.

Privalov, P. L.; O. B. Ptitsyn; Birshtei.Tm. Determination of stability of DNA double
helix in an aqueous medium, Biopolymers, 1969, 8, 559-571.

9.

Manning, G. S. Application of polyelectrolyte limiting laws to helix-coil transition of
DNA .1. Excess univalent cations, Biopolymers, 1972, 11, 937-947.

243

10.

Downey, C. D.; J. L. Fiore; C. D. Stoddard; J. H. Hodak; D. J. Nesbitt; A. Pardi. Metal
ion dependence, thermodynamics, and kinetics for intramolecular docking of a GAAA
tetraloop and receptor connected by a flexible linker, Biochemistry, 2006, 45, 3664–3673.

11.

Heilman-Miller, S. L.; J. Pan; D. Thirumalai; S. A. Woodson. Role of counterion
condensation in folding of the Tetrahymena ribozyme II. Counterion-dependence offolding kinetics, J. Mol. Biol., 2001, 309, 57–68.

12.

Heilman-Miller, S. L.; D. Thirumalai; S. A. Woodson. Role of counterion condensation
in folding of the Tetrahymena ribozyme. I. Equilibrium stabilization by cations, J. Mol.
Biol., 2001, 306, 1157–1166.

13.

Nixon, P. L.; D. P. Giedroc. Energetics of a strongly pH dependent RNA tertiary
structure in a frameshifting pseudoknot, J. Mol. Biol., 2000, 296, 659–671.

14.

Takamoto, K.; Q. He; S. Morris; M. R. Chance; M. Brenowitz. Monovalent cations
mediate formation of native tertiary structure of the Tetrahymena thermophila ribozyme,
Nat. Struct. Biol., 2002, 9, 928-933.

15.

Uchida, T.; K. Takamoto; Q. He; M. R. Chance; M. Brenowitz. Multiple monovalent iondependent pathways for the folding of the L-21 Tetrahymena thermophila ribozyme, J.
Mol. Biol., 2003, 328, 463-478.

16.

Ke, A.; F. Ding; J. D. Batchelor; J. A. Doudna. Structural roles of monovalent cations in
the HDV ribozyme, Structure, 2007, 15, 281-287.

17.

Jiang, Y. F.; M. Xiao; P. Yin; Y. Zhang. Monovalent cations use multiple mechanisms toresolve ribozyme misfolding, RNA, 2006, 12, 561-566.

18.

Lambert, D.; D. Leipply; R. Shiman; D. E. Draper. The influence of monovalent cation
size on the stability of RNA tertiary structures, J. Mol. Biol., 2009, 390, 791-804.

19.

Shiman, R.; D. E. Draper. Stabilization of RNA tertiary structure by monovalent cations,J. Mol. Biol., 2000, 302, 79–91.

20.

Uchida, T.; Q. He; C. Y. Ralston; M. Brenowitz; M. R. Chance. Linkage of monovalentand divalent ion binding in the folding of the P4-P6 domain of the Tetrahymena
ribozyme, Biochemistry, 2002, 41, 5799-5806.

21.

Basu, S.; R. P. Rambo; J. Strauss-Soukup; J. H. Cate; A. R. Ferre-D'Amare; S. A.
Strobel; J. A. Doudna. A specific monovalent metal ion integral to the AA platform of the
RNA tetraloop receptor, Nat. Struct. Biol., 1998, 5, 986–992.

22.

Wang, Y. X.; M. Lu; D. E. Draper. Specific ammonium ion requirement for functional
ribosomal-RNA tertiary structure, Biochemistry, 1993, 32, 12279-12282.

23.

Draper, D. E. RNA folding: Thermodynamic and molecular descriptions of the roles ofions, Biophys. J., 2008, 95, 5489-5495.

244

24.

Anderson, C. F.; M. T. Record. Salt nucleic-acid interactions, Annu. Rev. Phys. Chem.,
1995, 46, 657–700.

25.

Tinoco, I.; C. Bustamante. How RNA folds, J. Mol. Biol., 1999, 293, 271–281.

26.

Sosnick, T. R.; T. Pan. RNA folding: Models and perspectives, Curr. Opin. Struct. Biol.,
2003, 13, 309–316.

27.

Woese, C. R.; S. Winker; R. R. Gutell. Architecture of ribosomal-RNA: "Constraints on
the sequence of Tetraloops", Proc. Natl. Acad. Sci. USA, 1990, 87, 8467-8471.

28.

Hodak, J. H.; C. D. Downey; J. L. Fiore; A. Pardi; D. J. Nesbitt. Docking kinetics andequilibrium of a GAAA tetraloop-receptor motif probed by single-molecule FRET, Proc.
Natl. Acad. Sci. USA, 2005, 102, 10505–10510.

29.

Cate, J. H.; A. R. Gooding; E. Podell; K. H. Zhou; B. L. Golden; C. E. Kundrot; T. R.
Cech; J. A. Doudna. Crystal structure of a group I ribozyme domain: Principles of RNA
packing, Science, 1996, 273, 1678–1685.

30.

Batey, R. T.; R. P. Rambo; J. A. Doudna. Tertiary motifs in RNA structure and folding,Angew. Chem. Int. Ed., 1999, 38, 2327–2343.

31.

Costa, M.; F. Michel. Frequent use of the same tertiary motif by self-folding RNAs,
Embo. J., 1995, 14, 1276–1285.

32.

Vander Meulen, K. A.; J. H. Davis; T. R. Foster; T. Record; S. E. Butcher.
Thermodynamics and folding pathway of tetraloop receptor-mediated RNA helicalpacking, J. Mol. Biol., 2008, 384, 702-717.

33.

Young, B. T.; S. K. Silverman. The GAAA tetraloop-receptor interaction contributes
differentially to folding thermodynamics and kinetics for the P4-P6 RNA domain,
Biochemistry, 2002, 41, 12271–12276.

34.

Sattin, B. D.; W. Zhao; K. Travers; S. Chut; D. Herschlag. Direct measurement of tertiary
contact cooperativity in RNA folding, J. Am. Chem. Soc., 2008, 130, 6085–6087.

35.

Fiore, J. L.; J. H. Hodak; O. Piestert; C. D. Downey; D. J. Nesbitt. Monovalent andDivalent Promoted GAAA-Tetraloop–Receptor Tertiary Interactions from Freely
Diffusing Single-Molecule Studies, Biophys. J., 2008, 95, 3892-3905.

36.

Fiore, J. L.; B. Kraemer; F. Koberling; R. Erdmann; D. J. Nesbitt. Enthalpy-Driven RNA
Folding: Single-Molecule Thermodynamics of Tetraloop–Receptor Tertiary Interaction,
Biochemistry, 2009, 48, 2550–2558.

37.

Butcher, S. E.; T. Dieckmann; J. Feigon. Solution structure of a GAAA tetraloop receptor
RNA, Embo. J., 1997, 16, 7490–7499.

245

38.

Davis, J. H.; M. Tonelli; L. G. Scott; L. Jaeger; J. R. Williamson; S. E. Butcher. RNA
helical packing in solution: NMR structure of a 30 kDa GAAA tetraloop-receptor
complex J. Mol. Biol., 2005, 351, 371–382.

39.

Heus, H. A.; A. Pardi. Structural features that give rise to the unusual stability of RNA
hairpins containing GNRA loops, Science, 1991, 253, 191-194.

40.

Jucker, F. M.; H. A. Heus; P. F. Yip; E. H. M. Moors; A. Pardi. A network ofheterogeneous hydrogen bonds in GNRA tetraloops, J. Mol. Biol., 1996, 264, 968-980.

41.

Aleman, E. A.; R. Lamichhane; D. Rueda. Exploring RNA Folding One Molecule at aTime, Curr. Opin. Chem. Biol., 2008, 12, 647-654.

42.

Bokinsky, G.; X. W. Zhuang. Single-molecule RNA folding, Acc. Chem. Res., 2005, 38,
566–573.

43.

Blanco, M.; N. G. Walter. Analysis of complex single-molecule FRET time trajectories,
Methods Enzymol., 2010, 472, 153-178.

44.

Bartley, L. E.; X. W. Zhuang; R. Das; S. Chu; D. Herschlag. Exploration of theTransition State for Tertiary Structure Formation Between an RNA Helix and a Large
Structured RNA, J. Mol. Biol., 2003, 328, 1011–1026.

45.

Zhou, Y. J.; X. W. Zhuang. Robust reconstruction of the rate constant distribution using
the phase function method, Biophys. J., 2006, 91, 4045-4053.

46.

Leipply, D.; D. Lambert; D. E. Draper. Ion-RNA interactions: Thermodynamic analysisof the effects of mono- and divalent ions on RNA conformational equilibria, Methods
Enzymol., 2009, 469, 433-463.

47.

Bokinsky, G.; D. Rueda; V. K. Misra; M. M. Rhodes; A. Gordus; H. P. Babcock; N. G.
Walter; X. W. Zhuang. Single-molecule transition-state analysis of RNA folding, Proc.
Natl. Acad. Sci. USA, 2003, 100, 9302–9307.

48.

Fiore, J. L.; E. D. Holmstrom; D. J. Nesbitt. An Entropic origin of Mg2+-Facilitated RNA
Folding, Proc. Natl. Acad. Sci. USA, 2011, (Accepted).

49.

Hanggi, P.; P. Talkner; M. Borkovec. Reaction-rate theory - 50 years after Kramers, Rev.
Mod. Phys., 1990, 62, 251-341.

50.

Vander Meulen, K. A.; S. E. Butcher. Characterization of the kinetic and thermodynamic
landscape of RNA folding using a novel application of isothermal titration calorimetry,
2011, DOI:10.1093/nar/gkr894.

51.

Davis, J. H.; T. R. Foster; M. Tonelli; S. E. Butcher. Role of metal ions in the tetraloopreceptor complex as analyzed by NMR, RNA, 2007, 13, 76-86.

246

52.

Buchmueller, K. L.; A. E. Webb; D. A. Richardson; K. M. Weeks. A collapsed nonnative RNA folding state, Nat. Struct. Biol., 2000, 7, 362-366.

53.

Swisher, J. F.; L. H. J. Su; M. Brenowitz; V. E. Anderson; A. M. Pyle. Productive folding
to the native state by a group II intron ribozyme, J. Mol. Biol., 2002, 315, 297-310.

54.

Silverman, S. K.; T. R. Cech. An early transition state for folding of the P4-P6 RNA
domain, RNA, 2001, 7, 161–166.

55.

Pljevaljcic, G.; D. Klostermeier; D. P. Millar. The tertiary structure of the hairpin
ribozyme is formed through a slow conformational search, Biochemistry, 2005, 44, 48704876.

56.

Szewczak, A. A.; E. R. Podell; P. C. Bevilacqua; T. R. Cech. Thermodynamic stability of
the P4-P6 domain RNA tertiary structure measured by temperature gradient gelelectrophoresis, Biochemistry, 1998, 37, 11162–11170.

57.

Vander Meulen, K. A.; S. E. Butcher. Characterization of the kinetic and thermodynamic
landscape of RNA folding using a novel application of isothermal titration calorimetry,Nucl. Acids Res., 2012, 40, 2140-2151.

58.

Holbrook, J. A.; M. W. Capp; R. M. Saecker; M. T. Record. Enthalpy and heat capacitychanges for formation of an oligomeric DNA duplex: Interpretation in terms of coupledprocesses of formation and association of single-stranded helices, Biochemistry, 1999, 38,
8409-8422.

59.

Takach, J. C.; P. J. Mikulecky; A. L. Feig. Salt-dependent heat capacity changes for RNA
duplex formation, J. Am. Chem. Soc., 2004, 126, 6530-6531.

60.

Qin, P. Z.; J. Feigon; W. L. Hubbell. Site-directed spin labeling studies reveal solution
conformational changes in a GAAA tetraloop receptor upon Mg2+-dependent docking ofa GAAA tetraloop, J. Mol. Biol., 2005, 351, 1-8.

61.

Lambert, D.; D. Leipply; D. E. Draper. The osmolyte TMAO stabilizes native RNA
tertiary structures in the absence of Mg2+: Evidence for a large barrier to folding from
phosphate dehydration, J. Mol. Biol., 2010, 404, 138-157.

62.

Breslauer, K. J.; J. M. Sturtevant. Calorimetric investigation of single stranded basestacking in ribo-oligonucleotide-A7, Biophys. Chem., 1977, 7, 205-209.

63.

Seol, Y.; G. M. Skinner; K. Visscher. Elastic properties of a single-stranded charged
homopolymeric ribonucleotide, Phys. Rev. Lett., 2004, 93, 118102.

64.

Michelson, A. M.; C. Monny. Polynucleotides .8. Base stacking in polyuridylic acid,
Proc. Natl. Acad. Sci. U.S.A., 1966, 56, 1528-1534.

247

65.

Fiore, J. L.; E. D. Holmstrom; L. R. Feigland; J. H. Hodak; D. J. Nesbitt. The Role ofCounterion Valence and Size in the GAAA Tetraloop-Receptor Docking/Undocking
Kinetics, 2011, (In preperation).

66.

Holmstrom, E. D.; D. J. Nesbitt. Real-time infrared overtone laser control of temperaturein picoliter H2O samples: "Nanobathtubs" for single-molecule microscopy, J. Phys.
Chem. Lett., 2010, 1, 2264–2268.
Chapter 6:
Kinetic and Thermodynamic Origins of Osmolyte-Influenced Nucleic Acid Folding

1.

Draper, D. E. A guide to ions and RNA structure, RNA, 2004, 10, 335-343.

2.

Klein, D. J.; P. B. Moore; T. A. Steitz. The contribution of metal ions to the structural
stability of the large ribosomal subunit, RNA, 2004, 10, 1366-1379.

3.

Kurz, M. Compatible solute influence on nucleic acids: many questions but few answers,
Saline Systems, 2008, 4, 6.

4.

Yancey, P. H.; M. E. Clark; S. C. Hand; R. D. Bowlus; G. N. Somero. Living with water
stress: evolution of osmolyte systems, Science, 1982, 217, 1214-1222.

5.

Record, M. T., Jr.; E. S. Courtenay; D. S. Cayley; H. J. Guttman. Responses of E. coli toosmotic stress: large changes in amounts of cytoplasmic solutes and water, Trends
Biochem. Sci., 1998, 23, 143-148.

6.

Yancey, P. H.; G. N. Somero. Counteraction of urea destabilization of protein structure
by methylamine osmoregulatory compounds of elasmobranch fishes, Biochem. J., 1979,
183, 317-323.

7.

Record, M. T., Jr.; E. S. Courtenay; S. Cayley; H. J. Guttman. Biophysical compensation
mechanisms buffering E. coli protein-nucleic acid interactions against changing
environments, Trends Biochem. Sci., 1998, 23, 190-194.

8.

Liu, Y.; D. W. Bolen. The peptide backbone plays a dominant role in protein stabilization
by naturally occurring osmolytes, Biochemistry, 1995, 34, 12884-12891.

9.

Timasheff, S. N. Control of protein stability and reactions by weakly interacting
cosolvents: the simplicity of the complicated, Adv. Protein Chem., 1998, 51, 355-432.

10.

Courtenay, E. S.; M. W. Capp; R. M. Saecker; M. T. Record, Jr. Thermodynamic
analysis of interactions between denaturants and protein surface exposed on unfolding:interpretation of urea and guanidinium chloride m-values and their correlation withchanges in accessible surface area (ASA) using preferential interaction coefficients andthe local-bulk domain model, Proteins, 2000, 4, 72-85.

248

11.

Courtenay, E. S.; M. W. Capp; C. F. Anderson; M. T. Record, Jr. Vapor pressure
osmometry studies of osmolyte-protein interactions: implications for the action of
osmoprotectants in vivo and for the interpretation of "osmotic stress" experiments invitro, Biochemistry, 2000, 39, 4455-4471.

12.

Lambert, D.; D. E. Draper. Effects of Osmolytes on RNA Secondary and Tertiary
Structure Stabilities and RNA-Mg2+ Interactions, J. Mol. Biol., 2007, 370, 993-1005.

13.

Lambert, D.; D. Leipply; D. E. Draper. The osmolyte TMAO stabilizes native RNA
tertiary structures in the absence of Mg2+: Evidence for a large barrier to folding from
phosphate dehydration, J. Mol. Biol., 2010, 404, 138-157.

14.

Lambert, D.; D. E. Draper. Denaturation of RNA secondary and tertiary structure byurea: simple unfolded state models and free energy parameters account for measured mvalues, Biochemistry, 2012, 51, 9014-9026.

15.

Spink, C. H.; N. Garbett; J. B. Chaires. Enthalpies of DNA melting in the presence ofosmolytes, Biophys. Chem., 2007, 126, 176-185.

16.

Hong, J.; M. W. Capp; C. F. Anderson; R. M. Saecker; D. J. Felitsky; M. W. Anderson;
M. T. Record, Jr. Preferential interactions of glycine betaine and of urea with DNA:
implications for DNA hydration and for effects of these solutes on DNA stability,
Biochemistry, 2004, 43, 14744-14758.

17.

Felitsky, D. J.; J. G. Cannon; M. W. Capp; J. Hong; A. W. Van Wynsberghe; C. F.
Anderson; M. T. Record, Jr. The exclusion of glycine betaine from anionic biopolymer
surface: why glycine betaine is an effective osmoprotectant but also a compatible solute,
Biochemistry, 2004, 43, 14732-14743.

18.

Bennion, B. J.; V. Daggett. The molecular basis for the chemical denaturation of proteins
by urea, Proc. Natl. Acad. Sc.i USA, 2003, 100, 5142-5147.

19.

Greene, R. F., Jr.; C. N. Pace. Urea and guanidine hydrochloride denaturation ofribonuclease, lysozyme, alpha-chymotrypsin, and beta-lactoglobulin, J. Biol. Chem.,
1974, 249, 5388-5393.

20.

Tanford, C. Isothermal Unfolding of Globular Proteins in Aqueous Urea Solutions, J.
Am. Chem. Soc., 1964, 86, 2050-&.

21.

Shelton, V. M.; T. R. Sosnick; T. Pan. Applicability of Urea in the Thermodynamic
Analysis of Secondary and Tertiary RNA Folding, Biochemistry, 1999, 38, 16831-16839.

22.

Canchi, D. R.; A. E. Garcia. Cosolvent Effects on Protein Stability, Annu. Rev. Phys.
Chem., 2013.

23.

Schwinefus, J. J.; M. J. Kuprian; J. W. Lamppa; W. E. Merker; K. N. Dorn; G. W. Muth.
Human telomerase RNA pseudoknot and hairpin thermal stability with glycine betaine249and urea: Preferential interactions with RNA secondary and tertiary structures,
Biochemistry, 2007, 46, 9068-9079.
24.

Cho, S. S.; G. Reddy; J. E. Straub; D. Thirumalai. Entropic Stabilization of Proteins by
TMAO, J. Phys. Chem. B, 2011, 115, 13401-13407.

25.

Wang, A. J.; D. W. Bolen. A naturally occurring protective system in urea-rich cells:
Mechanism of osmolyte protection of proteins against urea denaturation, Biochemistry,
1997, 36, 9101-9108.

26.

Pincus, D. L.; C. Hyeon; D. Thirumalai. Effects of trimethylamine N-oxide (TMAO) andcrowding agents on the stability of RNA hairpins, J. Am. Chem. Soc., 2008, 130, 73647372.

27.

Priyakumar, U. D.; C. Hyeon; D. Thirumalai; A. D. MacKerell. Urea Destabilizes RNA
by Forming Stacking Interactions and Multiple Hydrogen Bonds with Nucleic Acid
Bases, J. Am. Chem. Soc., 2009, 131, 17759-17761.

28.

Tinoco, I.; R. L. Gonzalez. Biological mechanisms, one molecule at a time, Gene Dev.,
2011, 25, 1205-1231.

29.

Roy, R.; S. Hohng; T. Ha. A practical guide to single-molecule FRET, Nat. Methods,
2008, 5, 507-516.

30.

Li, P. T.; J. Vieregg; I. Tinoco, Jr. How RNA Unfolds and Refolds, Annu. Rev. Biochem.,
2008, 77, 77-100.

31.

Bokinsky, G.; X. Zhuang. Single-Molecule RNA Folding, Acc. Chem. Res., 2005, 38,
566-573.

32.

Aleman, E. A.; R. Lamichhane; D. Rueda. Exploring RNA folding one molecule at atime, Curr. Opin. Chem. Biol. 2008, 12, 647-654.

33.

Dupuis, N. F.; E. D. Holmstrom; D. J. Nesbitt. Single-Molecule Kinetics Reveal CationPromoted DNA Duplex Formation Through Ordering of Single-Stranded Helices,
Biophys. J., 2013, 105, 756-766.

34.

Downey, C. D.; J. L. Fiore; C. D. Stoddard; J. H. Hodak; D. J. Nesbitt; A. Pardi. Metal
ion dependence, thermodynamics, and kinetics for intramolecular docking of a GAAA
tetraloop and receptor connected by a flexible linker, Biochemistry, 2006, 45, 3664–3673.

35.

Fiore, J. L.; J. H. Hodak; O. Piestert; C. D. Downey; D. J. Nesbitt. Monovalent andDivalent Promoted GAAA-Tetraloop–Receptor Tertiary Interactions from Freely
Diffusing Single-Molecule Studies, Biophys. J., 2008, 95, 3892-3905.

36.

Fiore, J. L.; E. D. Holmstrom; L. R. Feigland; J. H. Hodak; D. J. Nesbitt. The Role ofCounterion Valence and Size in the GAAA Tetraloop-Receptor Docking/Undocking
Kinetics, 2012, J. Mol. Biol., 2012, 423, 198-216.

250

37.

Fiore, J. L.; E. D. Holmstrom; D. J. Nesbitt. Entropic origin of Mg2+-facilitated RNA
folding, Proc. Natl. Acad. Sci. USA, 2012, 109, 2902-2907.

38.

Fiore, J. L.; B. Kraemer; F. Koberling; R. Erdmann; D. J. Nesbitt. Enthalpy-Driven RNA
Folding: Single-Molecule Thermodynamics of Tetraloop–Receptor Tertiary Interaction,
Biochemistry, 2009, 48, 2550–2558.

39.

Hodak, J. H.; C. D. Downey; J. L. Fiore; A. Pardi; D. J. Nesbitt. Docking kinetics andequilibrium of a GAAA tetraloop-receptor motif probed by single-molecule FRET, Proc.
Natl. Acad. Sci. USA, 2005, 102, 10505–10510.

40.

Holmstrom, E. D.; J. L. Fiore; D. J. Nesbitt. Thermodynamic Origins of Monovalent
Facilitated RNA Folding, Biochemistry, 2012, 51, 3732-3743.

41.

Blanco, M.; N. G. Walter. Analysis of complex single-molecule FRET time trajectories,
Methods Enzymol., 2010, 472, 153-178.

42.

Bartley, L. E.; X. W. Zhuang; R. Das; S. Chu; D. Herschlag. Exploration of theTransition State for Tertiary Structure Formation Between an RNA Helix and a Large
Structured RNA, J. Mol. Biol., 2003, 328, 1011–1026.

43.

Zhou, Y. J.; X. W. Zhuang. Robust reconstruction of the rate constant distribution using
the phase function method, Biophys. J., 2006, 91, 4045-4053.

44.

Sclavi, B.; S. Woodson; M. Sullivan; M. R. Chance; M. Brenowitz. Time-resolved
synchrotron X-ray "footprinting", a new approach to the study of nucleic acid structureand function: application to protein-DNA interactions and RNA folding, J. Mol. Biol.,
1997, 266, 144-159.

45.

Pan, J.; D. Thirumalai; S. A. Woodson. Folding of RNA involves parallel pathways, J.
Mol. Biol., 1997, 273, 7-13.

46.

Pace, C. N. The stability of globular proteins, CRC Crit. Rev. Biochem., 1975, 3, 1-43.

47.

Myers, J. K.; C. N. Pace; J. M. Scholtz. Denaturant m values and heat capacity changes:
relation to changes in accessible surface areas of protein unfolding, Protein Sci., 1995, 4,
2138-2148.

48.

Fiore, J. L.; E. D. Holmstrom; D. J. Nesbitt. Entropic Origin of Mg2+-Facilitated RNA
Folding, Proc. Natl. Acad. Sci. USA, 2012, 109, 2902-2907.

49.

Holmstrom, E. D.; D. J. Nesbitt. Single-molecule fluorescence resonance energy transfer
studies of the human telomerase RNA pseudoknot: temperature-/urea-dependent folding
kinetics and thermodynamics, J. Phys. Chem. B, 2014, 118, 3853-3863.

50.

Dalgarno, P. A.; J. Bordello; R. Morris; P. St-Pierre; A. Dube; I. D. Samuel; D. A.
Lafontaine; J. C. Penedo. Single-molecule chemical denaturation of riboswitches, Nucleic
Acids Res., 2013, 41, 4253-4265.

251

51.

Dupuis, N. F.; E. D. Holmstrom; D. J. Nesbitt. Molecular-crowding effects on singlemolecule RNA folding/unfolding thermodynamics and kinetics, Proc. Natl. Acad. Sci.
USA, 2013. (in Press)

52.

Cornish-Bowden, A. Enthalpy-entropy compensation: a phantom phenomenon, J. Biosci.,
2002, 27, 121-126.

53.

Burg, M. B. Coordinate regulation of organic osmolytes in renal cells, Kidney Int., 1996,
49, 1684-1685.

54.

Aitken, C. E.; R. A. Marshall; J. D. Puglisi. An Oxygen Scavenging System for
Improvement of Dye Stability in Single-Molecule Fluorescence Experiments, Biophys. J.,
2008, 94, 1826-1835.
Chapter 7:
Single Molecule FRET Studies of the Human Telomerase RNA Pseudoknot:
Temperature/Urea Dependent Folding Kinetics and Thermodynamics

1.

Wynford-Thomas, D.; D. Kipling. The End-Replication Problem, Nature, 1997, 389,
551-551.

2.

Blackburn, E. H. Switching and Signaling at the Telomere, Cell, 2001, 106, 661-673.

3.

Blackburn, E. H. The End of the (DNA) Line, Nat. Struct. Biol., 2000, 7, 847-850.

4.

Greider, C. W.; E. H. Blackburn. Tracking Telomerase, Cell, 2004, 116, S83-86.

5.

Cech, T. R. Beginning to Understand the End of the Chromosome, Cell, 2004, 116, 273279.

6.

Hengesbach, M.; B. M. Akiyama; M. D. Stone. Single-Molecule Analysis of Telomerase
Structure and Function, Curr. Opin. Chem. Biol., 2011, 15, 845-852.

7.

Mihalusova, M.; J. Y. Wu; X. Zhuang. Functional Importance of Telomerase Pseudoknot
Revealed by Single-Molecule Analysis, Proc. Natl. Acad. Sci. USA, 2011, 108, 2033920344.

8.

Yeoman, J. A.; A. Orte; B. Ashbridge; D. Klenerman; S. Balasubramanian. RNA
Conformation in Catalytically Active Human Telomerase, J. Am. Chem. Soc., 2010, 132,2852-2853.

9.

Wu, J. Y.; M. D. Stone; X. Zhuang. A Single-Molecule Assay for Telomerase StructureFunction Analysis, Nucleic Acids Res., 2010, 38, e16.

10.

Gavory, G.; M. F. Symmons; Y. Krishnan Ghosh; D. Klenerman; S. Balasubramanian.
Structural Analysis of the Catalytic Core of Human Telomerase RNA by FRET and
Molecular Modeling, Biochemistry, 2006, 45, 13304-13311.

252

11.

Collins, K. The Biogenesis and Regulation of Telomerase Holoenzymes, Nat. Rev. Mol.
Cell. Biol., 2006, 7, 484-494.

12.

Chen, J. L.; C. W. Greider. Telomerase RNA Structure and Function: Implications for
Dyskeratosis Congenita, Trends Biochem. Sci., 2004, 29, 183-192.

13.

Chen, J. L.; M. A. Blasco; C. W. Greider. Secondary Structure of Vertebrate Telomerase
RNA, Cell, 2000, 100, 503-514.

14.

Gilley, D.; E. H. Blackburn. The Telomerase RNA Pseudoknot is Critical for the Stable
Assembly of a Catalytically Active Ribonucleoprotein, Proc. Natl. Acad. Sci. USA, 1999,
96, 6621-6625.

15.

Tzfati, Y.; Z. Knight; J. Roy; E. H. Blackburn. A Novel Pseudoknot Element is Essential
for the Action of a Yeast Telomerase, Genes Dev., 2003, 17, 1779-1788.

16.

Reipa, V.; G. Niaura; D. H. Atha. Conformational Analysis of the Telomerase RNA
Pseudoknot Hairpin by Raman Spectroscopy, RNA, 2007, 13, 108-115.

17.

Moriarty, T. J.; D. T. Marie-Egyptienne; C. Autexier. Functional Organization of Repeat
Addition Processivity and DNA Synthesis Determinants in the Human Telomerase
Multimer, Mol. Cell. Biol., 2004, 24, 3720-3733.

18.

Autexier, C.; R. Pruzan; W. D. Funk; C. W. Greider. Reconstitution of Human
Telomerase Activity and Identification of a Minimal Functional Region of the Human
Telomerase RNA, Embo. J., 1996, 15, 5928-5935.

19.

Bachand, F.; I. Triki; C. Autexier. Human Telomerase RNA-Protein Interactions, Nucleic
Acids Res., 2001, 29, 3385-3393.

20.

Kim, N. K.; Q. Zhang; J. Zhou; C. A. Theimer; R. D. Peterson; J. Feigon. Solution
Structure and Dynamics of the Wild-Type Pseudoknot of Human Telomerase RNA, J.
Mol. Biol., 2008, 384, 1249-1261.

21.

Comolli, L. R.; I. Smirnov; L. Xu; E. H. Blackburn; T. L. James. A Molecular Switch
Underlies a Human Telomerase Disease, Proc. Natl. Acad. Sci. USA, 2002, 99, 1699817003.

22.

Theimer, C. A.; C. A. Blois; J. Feigon. Structure of the Human Telomerase RNA
Pseudoknot Reveals Conserved Tertiary Interactions Essential for Function, Mol. Cell.,
2005, 17, 671-682.

23.

Theimer, C. A.; L. D. Finger; L. Trantirek; J. Feigon. Mutations Linked to Dyskeratosis
Congenita Cause Changes in the Structural Equilibrium in Telomerase RNA, Proc. Natl.
Acad. Sci. USA, 2003, 100, 449-454.

24.

Theimer, C. A.; L. D. Finger; J. Feigon. YNMG Tetraloop Formation by a Dyskeratosis
Congenita Mutation in Human Telomerase RNA, RNA, 2003, 9, 1446-1455.

253

25.

Chen, G.; J. D. Wen; I. Tinoco, Jr. Single-Molecule Mechanical Unfolding and Foldingof a Pseudoknot in Human Telomerase RNA, RNA, 2007, 13, 2175-2188.

26.

Hengesbach, M.; N. K. Kim; J. Feigon; M. D. Stone. Single-Molecule FRET Reveals the
Folding Dynamics of the Human Telomerase RNA Pseudoknot Domain, Angew. Chem.
Int. Ed. Engl., 2012, 51, 5876-5879.

27.

Cao, S.; S. J. Chen. Biphasic Folding Kinetics of RNA Pseudoknots and Telomerase
RNA Activity, J. Mol. Biol., 2007, 367, 909-924.

28.

Cao, S.; S. J. Chen. Predicting RNA Pseudoknot Folding Thermodynamics, Nucleic
Acids Res., 2006, 34, 2634-2652.

29.

Li, P. T.; J. Vieregg; I. Tinoco, Jr. How RNA Unfolds and Refolds, Annu. Rev. Biochem.,
2008, 77, 77-100.

30.

Bokinsky, G.; X. Zhuang. Single-Molecule RNA Folding, Acc. Chem. Res., 2005, 38,
566-573.

31.

Roy, R.; S. Hohng; T. Ha. A Practical Guide to Single-Molecule FRET, Nat. Methods,
2008, 5, 507-516.

32.

Zhao, R.; D. Rueda. RNA Folding Dynamics by Single-Molecule Fluorescence
Resonance Energy Transfer, Methods, 2009, 49, 112-117.

33.

Aleman, E. A.; R. Lamichhane; D. Rueda. Exploring RNA Folding One Molecule at aTime, Curr. Opin. Chem. Biol., 2008, 12, 647-654.

34.

Holmstrom, E. D.; J. L. Fiore; D. J. Nesbitt. Thermodynamic Origins of Monovalent
Facilitated RNA Folding, Biochemistry, 2012, 51, 3732-3743.

35.

Fiore, J. L.; E. D. Holmstrom; D. J. Nesbitt. Entropic Origin of Mg2+-Facilitated RNA
Folding, Proc. Natl. Acad. Sci. USA, 2012, 109, 2902-2907.

36.

Bartley, L. E.; X. W. Zhuang; R. Das; S. Chu; D. Herschlag. Exploration of theTransition State for Tertiary Structure Formation Between an RNA Helix and a Large
Structured RNA, J. Mol. Biol., 2003, 328, 1011–1026.

37.

Fiore, J. L.; B. Kraemer; F. Koberling; R. Erdmann; D. J. Nesbitt. Enthalpy-Driven RNA
Folding: Single-Molecule Thermodynamics of Tetraloop–Receptor Tertiary Interaction,
Biochemistry, 2009, 48, 2550–2558.

38.

Shelton, V. M.; T. R. Sosnick; T. Pan. Applicability of Urea in the Thermodynamic
Analysis of Secondary and Tertiary RNA Folding, Biochemistry, 1999, 38, 16831-16839.

39.

Lambert, D.; D. E. Draper. Effects of Osmolytes on RNA Secondary and Tertiary
Structure Stabilities and RNA-Mg2+ Interactions, J. Mol. Biol., 2007, 370, 993-1005.

254

40.

Fersht, A. R.; A. Matouschek; L. Serrano. The Folding of an Enzyme .1. Theory ofProtein Engineering Analysis of Stability and Pathway of Protein Folding, J. Mol. Biol.,
1992, 224, 771-782.

41.

Priyakumar, U. D.; C. Hyeon; D. Thirumalai; A. D. MacKerell. Urea Destabilizes RNA
by Forming Stacking Interactions and Multiple Hydrogen Bonds with Nucleic Acid
Bases, J. Am. Chem. Soc., 2009, 131, 17759-17761.

42.

Lambert, D.; D. E. Draper. Denaturation of RNA secondary and tertiary structure byurea: simple unfolded state models and free energy parameters account for measured mvalues, Biochemistry, 2012, 51, 9014-9026.

43.

Williams, S.; T. P. Causgrove; R. Gilmanshin; K. S. Fang; R. H. Callender; W. H.
Woodruff; R. B. Dyer. Fast Events in Protein Folding: Helix Melting and Formation in a
Small Peptide, Biochemistry, 1996, 35, 691-697.

44.

Yingling, Y. G.; B. A. Shapiro. The Impact of Dyskeratosis Congenita Mutations on the
Structure and Dynamics of the Human Telomerase RNA Pseudoknot Domain, J. Biomol.
Struct. Dyn., 2007, 24, 303-319.

45.

Akiyama, B. M.; M. D. Stone. Assembly of Complex RNAs by Splinted Ligation,
Methods Enzymol., 2009, 469, 27-46.

46.

Moore, M. J.; C. C. Query. Joining of RNAs by Splinted Ligation, Method Enzymol.,
2000, 317, 109-123.

47.

Stark, M. R.; J. A. Pleiss; M. Deras; S. A. Scaringe; S. D. Rader. An RNA LigaseMediated Method for the Efficient Creation of Large, Synthetic RNAs, RNA, 2006, 12,2014-2019.

48.

Moore, M. J., Query, C.C. Uses of Site-Specifically Modified RNAs Constructed by
RNA Ligation, In RNA:Protein Interactions: A Practical Approach (Smith, C. W. J.,
Ed.), pp 75-140, Oxford University Press Inc., New York.1998

49.

Chen, J. L.; C. W. Greider. Functional Analysis of the Pseudoknot Structure in Human
Telomerase RNA, Proc. Natl. Acad. Sci. USA, 2005, 102, 8080-8085; discussion 80778089.

50.

Lemay, J. F.; J. C. Penedo; R. Tremblay; D. M. Lilley; D. A. Lafontaine. Folding of the
Adenine Riboswitch, Chem. Biol., 2006, 13, 857-868.

51.

Lee, N. K.; A. N. Kapanidis; Y. Wang; X. Michalet; J. Mukhopadhyay; R. H. Ebright; S.
Weiss. Accurate FRET Measurements within Single Diffusing Biomolecules using
Alternating-Laser Excitation, Biophys. J., 2005, 88, 2939-2953.

52.

Kapanidis, A. N.; N. K. Lee; T. A. Laurence; S. Doose; E. Margeat; S. Weiss.
Fluorescence-Aided Molecule Sorting: Analysis of Structure and Interactions by255

Alternating-Laser Excitation of Single Molecules, Proc. Natl. Acad. Sci. USA, 2004, 101,
8936–8941.
53.

Kapanidis, A. N.; T. A. Laurence; N. K. Lee; E. Margeat; X. X. Kong; S. Weiss.
Alternating-Laser Excitation of Single Molecules, Acc. Chem. Res., 2005, 38, 523-533.

54.

Orte, A.; R. Clarke; S. Balasubramanian; D. Klenerman. Determination of the Fractionand Stoichiometry of Femtomolar Levels of Biomolecular Complexes in an Excess of
Monomer using Single-M olecule, Two-Color Coincidence Detection, Anal. Chem.,
2006, 78, 7707-7715.

55.

Fiore, J. L.; J. H. Hodak; O. Piestert; C. D. Downey; D. J. Nesbitt. Monovalent andDivalent Promoted GAAA-Tetraloop–Receptor Tertiary Interactions from Freely
Diffusing Single-Molecule Studies, Biophys. J., 2008, 95, 3892-3905.

56.

Aitken, C. E.; R. A. Marshall; J. D. Puglisi. An Oxygen Scavenging System for
Improvement of Dye Stability in Single-Molecule Fluorescence Experiments, Biophys. J.,
2008, 94, 1826-1835.
Chapter 8:
Vitamin B12-facilitates formation of a regulatory RNA kissing loop

1.

Tucker, B. J.; R. R. Breaker. Riboswitches as versatile gene control elements, Curr. Opin.
Struct. Biol., 2005, 15, 342-348.

2.

Nahvi, A.; N. Sudarsan; M. S. Ebert; X. Zou; K. L. Brown; R. R. Breaker. Genetic
control by a metabolite binding mRNA, Chem. Biol., 2002, 9, 1043.

3.

Nou, X.; R. J. Kadner. Adenosylcobalamin inhibits ribosome binding to btuB RNA,
Proc. Natl. Acad. Sci. USA, 2000, 97, 7190-7195.

4.

Banerjee, R.; S. W. Ragsdale. The many faces of vitamin B12: catalysis by cobalamindependent enzymes, Annu. Rev. Biochem., 2003, 72, 209-247.

5.

Nahvi, A.; J. E. Barrick; R. R. Breaker. Coenzyme B12 riboswitches are widespread
genetic control elements in prokaryotes, Nucleic Acids Res., 2004, 32, 143-150.

6.

Rodionov, D. A.; A. G. Vitreschak; A. A. Mironov; M. S. Gelfand. Comparative
genomics of the vitamin B12 metabolism and regulation in prokaryotes, J. Biol. Chem.,
2003, 278, 41148-41159.

7.

Martens, J. H.; H. Barg; M. J. Warren; D. Jahn. Microbial production of vitamin B12,
Appl. Microbiol. Biotechnol., 2002, 58, 275-285.

8.

Roth, J. R.; J. G. Lawrence; T. A. Bobik. Cobalamin (coenzyme B12): synthesis andbiological significance, Annu. Rev. Microbiol., 1996, 50, 137-181.

256

9.

Breaker, R. R. Riboswitches and the RNA world, Cold Spring Harb. Perspect. Biol.,2012, 4.

10.

Garst, A. D.; A. L. Edwards; R. T. Batey. Riboswitches: structures and mechanisms, Cold
Spring Harb. Perspect. Biol., 2011, 3.

11.

Roy, R.; S. Hohng; T. Ha. A Practical Guide to Single-Molecule FRET, Nat. Methods,
2008, 5, 507-516.

12.

Lemay, J. F.; J. C. Penedo; R. Tremblay; D. M. Lilley; D. A. Lafontaine. Folding of the
Adenine Riboswitch, Chem. Biol., 2006, 13, 857-868.

13.

Suddala, K. C.; A. J. Rinaldi; J. Feng; A. M. Mustoe; C. D. Eichhorn; J. A. Liberman; J.
E. Wedekind; H. M. Al-Hashimi; C. L. Brooks, 3rd; N. G. Walter. Single transcriptional
and translational preQ1 riboswitches adopt similar pre-folded ensembles that follow
distinct folding pathways into the same ligand-bound structure, Nucleic Acids Res., 2013,
41, 10462-10475.

14.

Souliere, M. F.; R. B. Altman; V. Schwarz; A. Haller; S. C. Blanchard; R. Micura.
Tuning a riboswitch response through structural extension of a pseudoknot, Proc. Natl.
Acad. Sci. USA, 2013, 110, E3256-3264.

15.

Haller, A.; R. B. Altman; M. F. Souliere; S. C. Blanchard; R. Micura. Folding and ligand
recognition of the TPP riboswitch aptamer at single-molecule resolution, Proc. Natl.
Acad. Sci. USA, 2013, 110, 4188-4193.

16.

Haller, A.; U. Rieder; M. Aigner; S. C. Blanchard; R. Micura. Conformational capture of
the SAM-II riboswitch, Nat. Chem. Biol., 2011, 7, 393-400.

17.

Fiegland, L. R.; A. D. Garst; R. T. Batey; D. J. Nesbitt. Single-molecule studies of thelysine riboswitch reveal effector-dependent conformational dynamics of the aptamer
domain, Biochemistry, 2012, 51, 9223-9233.

18.

Johnson, J. E., Jr.; F. E. Reyes; J. T. Polaski; R. T. Batey. B12 cofactors directly stabilize
an mRNA regulatory switch, Nature, 2012, 492, 133-137.

19.

Weinberg, Z.; J. X. Wang; J. Bogue; J. Yang; K. Corbino; R. H. Moy; R. R. Breaker.
Comparative genomics reveals 104 candidate structured RNAs from bacteria, archaea,
and their metagenomes, Genome Biol., 2010, 11, R31.

20.

Griffiths-Jones, S.; A. Bateman; M. Marshall; A. Khanna; S. R. Eddy. Rfam: an RNA
family database, Nucleic Acids Res., 2003, 31, 439-441.

21.

Xu, H.; Y. Li; C. Liu; Q. Wu; Y. Zhao; L. Lu; H. Tang. Fluorescence resonance energy
transfer between acridine orange and rhodamine 6G and its analytical application forvitamin B12 with flow-injection laser-induced fluorescence detection, Talanta, 2008, 77,
176-181.

257

22.

Ennifar, E.; P. Walter; B. Ehresmann; C. Ehresmann; P. Dumas. Crystal structures of
coaxially stacked kissing complexes of the HIV-1 RNA dimerization initiation site, Nat.
Struct. Biol., 2001, 8, 1064-1068.

23.

Horiya, S.; X. Li; G. Kawai; R. Saito; A. Katoh; K. Kobayashi; K. Harada. RNA LEGO:
magnesium-dependent formation of specific RNA assemblies through kissing
interactions, Chem. Biol., 2003, 10, 645-654.

24.

Tyrrell, J.; J. L. McGinnis; K. M. Weeks; G. J. Pielak. The cellular environment
stabilizes adenine riboswitch RNA structure, Biochemistry, 2013, 52, 8777-8785.

25.

Bronson, J. E.; J. Fei; J. M. Hofman; R. L. Gonzalez, Jr.; C. H. Wiggins. Learning rates
and states from biophysical time series: a Bayesian approach to model selection andsingle-molecule FRET data, Biophys. J., 2009, 97, 3196-3205.

26.

Andersen, A. A.; R. A. Collins. Intramolecular secondary structure rearrangement by the
kissing interaction of the Neurospora VS ribozyme, Proc. Natl. Acad. Sci. USA, 2001, 98,
7730-7735.

27.

Toor, N.; K. S. Keating; O. Fedorova; K. Rajashankar; J. Wang; A. M. Pyle. Tertiaryarchitecture of the Oceanobacillus iheyensis group II intron, RNA, 2010, 16, 57-69.

28.

Brunel, C.; R. Marquet; P. Romby; C. Ehresmann. RNA loop-loop interactions as
dynamic functional motifs, Biochimie, 2002, 84, 925-944.

29.

Wagner, E. G.; S. Brantl. Kissing and RNA stability in antisense control of plasmid
replication, Trends Biochem. Sci., 1998, 23, 451-454.

30.

Eguchi, Y.; J. Tomizawa. Complex formed by complementary RNA stem-loops and its
stabilization by a protein: function of CoIE1 Rom protein, Cell, 1990, 60, 199-209.

31.

Helmer-Citterich, M.; M. M. Anceschi; D. W. Banner; G. Cesareni. Control of ColE1
replication: low affinity specific binding of Rop (Rom) to RNAI and RNAII, Embo. J.,
1988, 7, 557-566.

32.

Di Primo, C. Real time analysis of the RNAI-RNAII-Rop complex by surface plasmon
resonance: from a decaying surface to a standard kinetic analysis, J. Mol. Recognit.,
2008, 21, 37-45.

33.

Banner, D. W.; M. Kokkinidis; D. Tsernoglou. Structure of the ColE1 rop protein at 1.7
A resolution, J. Mol. Biol., 1987, 196, 657-675.

34.

Williamson, J. R. Induced fit in RNA-protein recognition, Nat. Struct. Biol., 2000, 7,
834-837.

35.

Noeske, J.; J. Buck; B. Furtig; H. R. Nasiri; H. Schwalbe; J. Wohnert. Interplay of'induced fit' and preorganization in the ligand induced folding of the aptamer domain of
the guanine binding riboswitch, Nucleic Acids Res., 2007, 35, 572-583.

258

36.

Serganov, A.; Y. R. Yuan; O. Pikovskaya; A. Polonskaia; L. Malinina; A. T. Phan; C.
Hobartner; R. Micura; R. R. Breaker; D. J. Patel. Structural basis for discriminative
regulation of gene expression by adenine- and guanine-sensing mRNAs, Chem. Biol.,
2004, 11, 1729-1741.

37.

Zhang, J.; M. W. Lau; A. R. Ferre-D'Amare. Ribozymes and riboswitches: modulation of
RNA function by small molecules, Biochemistry, 2010, 49, 9123-9131.

38.

Bernstein, J. A.; A. B. Khodursky; P. H. Lin; S. Lin-Chao; S. N. Cohen. Global analysis
of mRNA decay and abundance in Escherichia coli at single-gene resolution using twocolor fluorescent DNA microarrays, Proc. Natl. Acad. Sci USA, 2002, 99, 9697-9702.

39.

Moore, M. J., Query, C.C. Uses of Site-Specifically Modified RNAs Constructed by
RNA Ligation, In RNA:Protein Interactions: A Practical Approach (Smith, C. W. J.,
Ed.), pp 75-140, Oxford University Press Inc., New York.1998

40.

Proudnikov, D.; A. Mirzabekov. Chemical methods of DNA and RNA fluorescentlabeling, Nucleic Acids Res., 1996, 24, 4535-4542.

41.

Kapanidis, A. N.; T. A. Laurence; N. K. Lee; E. Margeat; X. Kong; S. Weiss.
Alternating-laser excitation of single molecules, Acc. Chem. Res., 2005, 38, 523-533.

42.

Aitken, C. E.; R. A. Marshall; J. D. Puglisi. An Oxygen Scavenging System for
Improvement of Dye Stability in Single-Molecule Fluorescence Experiments, Biophys. J.,
2008, 94, 1826-1835.

43.

Hodak, J. H.; C. D. Downey; J. L. Fiore; A. Pardi; D. J. Nesbitt. Docking kinetics andequilibrium of a GAAA tetraloop-receptor motif probed by single-molecule FRET, Proc.
Natl. Acad. Sci. USA, 2005, 102, 10505–10510.

44.

Baba, T.; T. Ara; M. Hasegawa; Y. Takai; Y. Okumura; M. Baba; K. A. Datsenko; M.
Tomita; B. L. Wanner; H. Mori. Construction of Escherichia coli K-12 in-frame, singlegene knockout mutants: the Keio collection, Mol. Syst. Biol., 2006, 2, 2006 0008.

259