LIGHTING UP LIFE: FLUORESCENCE STUDIES OF THE SHAKER K+

CHANNEL AND MYOSIN V IN ACTION

BY

GREGORY E. SNYDER

B.S., University of Texas, 1995B.A., University of Texas, 1995

THESIS

Submitted in partial fulfillment of the requirementsfor the degree of Doctor of Philosophy in Physics

in the Graduate College of theUniversity of Illinois at Urbana-Champaign, 2003

Urbana, Illinois

c© Copyright by Gregory E. Snyder, 2003

LIGHTING UP LIFE: FLUORESCENCE STUDIES OF THE SHAKER K+

CHANNEL AND MYOSIN V IN ACTION

Gregory E. Snyder, Ph.D.Department of Physics

University of Illinois at Urbana-Champaign, 2003Paul R. Selvin, Advisor

Fluorescence spectroscopy provides a wide variety of techniques for studying the

conformational changes which proteins undergo as they function. In this thesis I

present experiments on two different proteins using a variety of fluorescence tech-

niques. One protein is the Shaker potassium channel, a protein which contributes to

the action potential in neurons by controlling the flow of K+ across the cell mem-

brane. The other is myosin V, a biomolecular motor involved in intracellular cargo

transport. The fluorescence techniques include LRET, FRAP, and FIONA, whose

meanings I describe immediately below.

Fluorescence Resonance Energy Transfer (FRET) is a technique for measuring

distances (2–10nm) shorter than the wavelength of light (∼500nm) by exploiting theability of two fluorescent dyes to transfer energy via interaction of their transition

dipole moments with an efficiency that depends on their separation. Lanthanide-

based Resonance Energy Transfer (LRET) is a modification of the technique in which

a luminescent lanthanide chelate, such as Tb3+-DTPA-cs124, is used in place of a con-

ventional organic fluorophore. LRET has several advantages over conventional FRET,

including a lower sensitivity to the relative orientation of the dipoles of the two dyes.

In order to verify the relative insensitivity of LRET to the orientation of the dyes,

the anisotropy of two lanthanide chelates, Tb3+-DTPA-cs124 and Eu3+-DTPA-cs124

was measured. The emission of Tb3+-DTPA-cs124 is found to be unpolarized, but the

emission of Eu3+-DTPA-cs124 is found to have a wavelength-dependent polarization.

The first protein I studied is the Shaker potassium channel, a voltage gated ion

channel selective for potassium ions. For many years it has been known that a par-

ticular region of the protein, called S4, contains a large number of positive charges

at physiological pH and moves in response to changes in the electrostatic potential

across the cell membrane. The motion of S4 is coupled to the “gate” which opens and

closes the pore. We used LRET to measure distances between identical sites on the

four subunits of Shaker and find no evidence for the traditional model of S4 motion, a

large translation across the membrane. Rather, our results suggest a model in which

S4 undergoes the more subtle motion of a rotation and possibly a tilt.

iii

Since LRET is fundamentally capable of measuring only linear distances, we

wished to detect the rotation of S4 directly to verify our model. However, if the

entire channel were free to rotate in the membrane on a time scale comparable or

faster than the putative S4 rotation, ∼10 ms, it would be impossible to observe S4rotation. As a first control, we measured translational diffusion of the channel to see

if it was free to move in the membrane or anchored in some fashion. In addition, the

question of how the channels are localized is itself of biological importance. An in-

strument to conduct experiments using Fluorescence Recovery After Photobleaching

(FRAP) was constructed. The experiments revealed the channel to be translationally

immobile. Further experiments were performed to identify the cause of its immo-

bility. Possibilities we investigated include attachment to cytoskeletal elements and

sequestration in lipid rafts. Disruption of the cytoskeleton by chemical means and

removal of putative binding sites from the protein did not free it to start diffusing.

However, we found that increasing doses of dimethyl sulfoxide (DMSO) could produce

increasing channel mobility by a non-specific mechanism. Preliminary measurements

to test an apparatus to detect the rotation are also discussed.

The other protein I studied is myosin V which walks bipedally along actin filaments

taking steps which displace the center of mass 37 nm per step. A controversy over

the nature of this movement was settled earlier this year by other members of our

laboratory when they showed that myosin V processes in a hand-over-hand manner,

with each of the heads alternating in the leading and trailing positions. To do this,

they developed a new technique which they called FIONA for Fluorescence Imaging

with One Nanometer Accuracy which allowed them to localize individual molecules of

myosin V to within 1.5nm on the surface of a microscope cover slip. I have extended

and applied FIONA to a mutant of myosin V with a molecule of the enhanced Green

Fluorescent Protein (eGFP) fused to one head of the dimer. I demonstrated that

eGFP is suitable to use with FIONA, confirm the hand-over-hand model of myosin

procession, and argue that in aggregate, the data on myosin’s steps from our lab

implies that myosin V adopts an asymmetric, “telemark skier,” configuration between

steps.

iv

For Tania and all of my parents

v

Acknowledgments

I have benefitted from my interaction with many people over the past seven years,

and to them I owe gratitude and humble thanks. Some of them are:

• My parents, for a lifetime of encouragement and support.

• Tania Chakrabarty, for a collaboration that extends beyond the lab, for dedi-cation and sacrifice I don’t deserve, without whom I probably would still be in

grad school. You give me a reason and desire to always improve.

• My advisor, Professor Paul Selvin, who made all of this possible, who taughtme patience, persistence, to discern what’s important, and to remain focused on

goals, whose optimism and enthusiasm kept me returning to a dark room with

green and blue lights night after night, and whose genius for obtaining funding

kept the toys rolling in.

• Jim Sellers (NIH) for providing eGFP-myosin samples on a moment’s noticeand his post-doc, Takeshi Sakamoto, for letting me use them.

• Professor Francisco Bezanilla (UCLA), for sharing his encyclopedic knowledgeof electronics and ion channels and for hosting my visit to his lab.

• Professor Enrico Gratton for allowing me to use his laser and take up space inhis lab for two years, and Doctor Theodore Hazlett for getting me started in

fluorescence, spending so many hours with me on L3 and PC2, and for always

having the time to answer questions.

• Professor David Piston (Vanderbilt) for donating the plasmid for eGFP.

• Professor Michael Weissman for giving a productive start to my tenure in thePhysics Department.

• Professors Michel Bellini and Philip Best for instruction and the use of equip-ment.

vi

• Matthew Gordon for singing at my wedding and keeping me company duringthose long nights in the LFD. Thanks for all the extra awesomeness in the

FRAP apparatus.

• Ahmet Yildiz for helping me to get started and keep going with FIONA.

• David Posson for consultation and companionship.

• The rest of the Selvin lab, Pinghua Ge, Jiyan Chen, Anne Gershenson, EvanGraves, Comert Kural, Jeff Reifenberger, Hyokeun Park, and Ming Xiao, for

their support and assistance.

• Members of the Bezanilla lab, Albert Cha, Dorine Starace, and Kwame Asamoah,for teaching me voltage clamping, spending hours on the phone to discussing

experiments, and for the additional hours spent on preparing samples.

• Members of the LFD, especially Don Lamb for his patience, assistance, anduseful discussions, Nicholas Barry for lending his expertise in optics, Jochen

Mueller and Yan Chen for their advice, and Osman Akcakir for the extra com-

pany on the night shift and for easing the transition to Ann Arbor for Tania

and me.

• Kevin O’Brien, Robert Merithew, and Michael Rabin for their help in the Weiss-man lab.

• Justin Gullingsrud, the C-dog, for being a companion, competitor, and unrealchess playa. Together we trained for four marathons, ran two of them, and

managed to remain in sync for five years of grad school. You’re the best, man.

• Erich Mueller and Ben McMahon for their incurable optimism and for havingso many answers.

• My first year study partners without whom I could not have passed 462 orthe qual, and who gave their support throughout the many years: Bill Neils,

Kevin Paul, Vishnu Jejjala, and Tony Bonetti. Together we made a pretty good

physicist.

• Vishnu Jejjala and Christopher Bouchard for believing I could be a theorist.

• Professor Dan Eads (Albert Einstein College of Medicine) for explaining at thevery beginning that a Ph.D. is seldom obtained alone. Looks like you were

right.

vii

In addition, I am grateful for the financial support of the Carver Trust Foundation,

NSF DBI-02-9984841, NIH AR44420, and the Molecular Biophysics Training Grant

PHS T32 GM08276.

viii

Contents

Chapter

1 Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1

1.1 Fluorescence spectroscopy . . . . . . . . . . . . . . . . . . . . . . . . 1

1.2 The Shaker potassium channel . . . . . . . . . . . . . . . . . . . . . . 3

1.2.1 The general structure of Shaker . . . . . . . . . . . . . . . . . 5

1.2.2 The voltage sensor . . . . . . . . . . . . . . . . . . . . . . . . 7

1.3 Single molecule spectroscopy and myosin V . . . . . . . . . . . . . . . 9

1.3.1 The green fluorescent protein . . . . . . . . . . . . . . . . . . 10

1.3.2 Myosin V . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12

2 Applications of luminescent lanthanide chelates . . . . . . . . . . . . . . . . . . . 19

2.1 Fluorescence resonance energy transfer

(FRET) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19

2.2 Lanthanide-based resonance energy transfer

(LRET) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23

2.3 Anisotropy of lanthanide chelates . . . . . . . . . . . . . . . . . . . . 25

3 The motion of the voltage sensor of Shaker detected by LRET . . . . 29

3.1 Materials and methods . . . . . . . . . . . . . . . . . . . . . . . . . . 29

3.1.1 Sample preparation . . . . . . . . . . . . . . . . . . . . . . . . 29

3.1.2 Labeling with fluorescent dyes . . . . . . . . . . . . . . . . . . 30

3.1.3 Two electrode voltage clamp . . . . . . . . . . . . . . . . . . . 31

3.1.4 Fluorescence detection . . . . . . . . . . . . . . . . . . . . . . 32

3.2 Results and discussion . . . . . . . . . . . . . . . . . . . . . . . . . . 32

3.3 The Big Picture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 37

4 Mobility of Shaker in the oocyte membrane . . . . . . . . . . . . . . . . . . . . . . 39

4.1 Fluorescence Recovery After Photobleaching (FRAP) . . . . . . . . . 40

4.2 Materials and methods . . . . . . . . . . . . . . . . . . . . . . . . . . 41

ix

4.2.1 Sample preparation . . . . . . . . . . . . . . . . . . . . . . . . 41

4.2.2 Confocal microscopy . . . . . . . . . . . . . . . . . . . . . . . 42

4.3 Results and discussion . . . . . . . . . . . . . . . . . . . . . . . . . . 44

4.3.1 EGFP-Shaker . . . . . . . . . . . . . . . . . . . . . . . . . . . 44

4.3.2 TMR-Shaker and attachment to the cytoskeleton . . . . . . . 45

4.4 Polarized Fluorescence Recovery After Photobleaching (pFRAP) . . . 51

5 Myosin V . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 56

5.1 Materials and methods . . . . . . . . . . . . . . . . . . . . . . . . . . 56

5.1.1 Total internal reflection fluorescence microscopy (TIRF) . . . 56

5.1.2 General sample chamber preparation . . . . . . . . . . . . . . 58

5.1.3 Biotinylating eGFP . . . . . . . . . . . . . . . . . . . . . . . . 59

5.1.4 Sample chamber preparation for stage stepping experiment . . 60

5.1.5 Myosin preparation . . . . . . . . . . . . . . . . . . . . . . . . 60

5.1.6 Sample chamber preparation for myosin V step size measurement 61

5.1.7 Data acquisition and analysis . . . . . . . . . . . . . . . . . . 62

5.1.8 Theoretical considerations . . . . . . . . . . . . . . . . . . . . 62

5.2 Results and discussion . . . . . . . . . . . . . . . . . . . . . . . . . . 64

5.2.1 Control experiments . . . . . . . . . . . . . . . . . . . . . . . 64

5.2.2 Measuring eGFP-myosin V step sizes . . . . . . . . . . . . . . 65

5.2.3 Using eGFP with FIONA . . . . . . . . . . . . . . . . . . . . 69

5.2.4 Telemark skier: Position and nature of the bend . . . . . . . . 71

5.3 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 73

6 Future directions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 74

6.1 Myosin V . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 74

6.2 E2GFP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 76

A The meaning of mutation notation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 77

B Protocols for preparing and handling Xenopus oocytes . . . . . . . . . . . 78

B.1 Solutions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 78

B.2 Mutagenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79

B.3 Oocyte preparation . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79

B.4 Injection with mRNA . . . . . . . . . . . . . . . . . . . . . . . . . . . 80

B.5 Pre-blocking . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 81

Vita. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91

x

Chapter 1

Introduction

1.1 Fluorescence spectroscopy

Fluorescence spectroscopy is one of the most powerful tools in structural biology. It

encompasses a broad range of techniques for exploring biological molecules in cuvettes

or in cells, in ensembles or individually.

Fluorescence is the process of light emission which occurs when a molecule absorbs

a photon, is excited to a singlet state, and emits a photon of lower energy (longer

wavelength). Figure 1.1 is a simplified diagram of the transitions between energy

levels in a fluorescing molecule, commonly called a JabÃloński diagram after Alexander

JabÃloński who first used an energy level diagram to describe fluorescence [1] & [2].

When a fluorescent molecule1 absorbs a photon (hν in Figure 1.1, left), it is

excited from the singlet ground state, S0, to a vibrational sub-state of one of the

singlet excited states, S1, S2, etc.). Absorption occurs in ∼1 fs. The large numberof closely spaced vibrational sub-states causes the absorption spectra of fluorophores

to appear continuous over a range of wavelengths (Figure 1.1, right). In contrast,

the absorption spectra of atoms are discrete because they have no internal structure

1I will use the terms “dye,” “fluorophore,” and “fluorescent molecule” interchangeably in thecoming chapters.

1

400 500 600 700wavelength (nm)

fluo

resc

ence

inte

nsity

(a.

u.)

absorptionemission

Figure 1.1: Left A JabÃloński diagram. Right Absorption and emission spectra of tetram-ethylrhodamine [3].

capable of dissipating energy. A short time (∼1 ps) after absorption, the moleculerelaxes to the lowest-energy vibrational sub-state of S1 by “internal conversion” of

the energy to heat. The molecule typically spends 1–10ns in this state, a time simply

referred to as the excited state lifetime, or the fluorescence lifetime. The molecule

then usually de-excites to one of the vibrational sub-states of S0 by emitting a photon.

The large number of closely spaced vibrational sub-states of S0 gives the fluorescence

emission spectrum its smoothness and breadth. The emission spectrum is redder than

the absorption spectrum because of the energy lost to internal conversion.

A fluorescent molecule may also relax from the excited state to a triplet state (T1)

by a process called “intersystem crossing.” Light emission from the triplet state is

known as “phosphorescence.” Because T1 is usually of lower energy than S1, phos-

phorescence is redder than fluorescence. The lifetime of T1 (∼1 ms−1) is also longerthan the fluorescence lifetime because it requires the excited electron to undergo a

spin flip, which only occurs through the weak spin-orbit coupling. Phosphorescence

is not observed in most experiments because collisions between the fluorophore and

other molecules in solution, particularly O2, can occur during the long lifetime of T1

and enable the molecule to de-excite without emitting a photon.

Flourescence enables the experimentalist to use optics to detect structural changes

in proteins while circumventing fundamental limits on optical resolution. According to

2

the Rayleigh criterion, in order for two structures to be resolved by a light microscope,

they must be separated by a distance roughly half the wavelength of the light used

to probe them. For optical wavelengths, this comes out to about 250 nm. The size

of proteins, however, is on the order of nanometers, making it impossible to directly

visualize them, let alone their internal structures.

Fluorescence has the advantage over other biophysical techniques of requiring

small quantities of the protein being studied. Most spectroscopic techniques used to

probe biological molecules such as nuclear magnetic resonance, electron paramagnetic

resonance, and x-ray crystallography use concentrations ranging from micromolar to

millimolar which involves purification of several milligrams of protein. Fluorescence

spectroscopy is sensitive enough to work on sub-nanomolar sample concentrations

and even on single molecules. Fluorescent labels are also more sensitive and safer to

use than radioactive labels. Fluorescence takes advantage of optical technology which

has been in development for centuries, but modern advances in light sources, lenses,

filters, and detectors were needed to fully realize its capabilities.

In the Selvin laboratory, we use fluorescence to study dynamic properties of pro-

teins. The two proteins I have studied are very different. The first is the Shaker

potassium channel which is a model system for the study of voltage gated ion chan-

nels, the proteins responsible for electrical signaling in neurons and muscle. The other

is myosin V, a molecular motor responsible for moving cargo around in the cell. I

will turn my attention now to introducing these proteins, and introduce the specific

fluorescence techniques I used in later chapters as they are required.

1.2 The Shaker potassium channel

The first biological system I studied was the Shaker voltage gated K+ channel. This

section contains background about the biology of ion channels and the structure of

Shaker in order to motivate the research discussed in Chapter 3.

3

Ion channels are proteins which reside in cell membranes in order to provide

ions with a hydrophilic passage through the hydrophobic membrane. Voltage gated

ion channels are ion channels which open and close with a probability biased by

the electrostatic potential across the cell membrane. They provide the cell with a

mechanism for regulating its membrane potential and are important for signaling in

muscles and neurons.

The Shaker potassium channel was the first voltage gated potassium channel to

be cloned [4] and has served as a model system for understanding potassium channels

in specific and voltage gated channels in general because of its high expression in non-

native expression systems (cf. Section 3.1.1). The Shaker channel comes from the

fruit fly, Drosophila melanogaster, and is so named because mutations in the channel

cause the flies to shake. Drosophila has provided us with examples of the three

other classes of potassium channels, Shal, Shab, and Shaw, and all known potassium

channels are classified by their similarity to one of the Drosophila types.

Potassium channels are so named because they are more permeable to potassium

than to other monovalent cations, particularly sodium. A channel’s permeability for

potassium may be as much as 104 higher than for sodium, but the selectivity of Shaker

is closer to 50-fold [5]. Despite this high selectivity, the channel allows ions to flow

across the membrane at a rate close to the diffusion limit [6].

Since 1991, when mutations in an ion channel were first linked to a disease2, ion

channels have been implicated in an ever increasing number of disorders. Some of

the symptoms of ion channel diseases include muscle spasms, fits of muscle weakness,

epileptic seizures, cardiac arrhythmias, and migraine headaches [7].

2Mutations in an adult muscle sodium channel were found to lead to hyperkalemic periodicparalysis, a rare disease characterized by attacks of muscle weakness and associated with elevatedlevels of K+ in the blood.

4

CLOSED

OPEN

S5

OUTSIDE

INSIDE

S4

S6S5S1

S3S2

S6

S4

S3

S2

K374

E293

D316

E283

R362

R365

R368

R371

INSIDE

S5

OUTSIDES4

S6S5S1

S3S2

S6

S4

S3

S2

intracellular

vestibule

extracellular

vestibule

S4

Figure 1.2: Left Idealization of the structure of Shaker compiled from crystal structures.Figure taken from [8]. Right A model of the arrangement of the transmembrane segmentsof Shaker which accounts for the putative rotation of S4 based on the work presented inChapter 3

1.2.1 The general structure of Shaker

Shaker is a tetramer of identical subunits surrounding a central pore. Each subunit

consists of a chain of 656 amino acids which forms several distinct domains. The

crystal structure of Shaker has not been solved, but the left side of Figure 1.2 shows

a cartoon assembled from pieces of structures from homologous proteins.

Until five years ago, little was known about the arrangement of the protein in the

membrane. Based largely on refinements of the original hydropathy3 analysis made

when the gene for Shaker was first cloned [4], the transmembrane region was believed

to consist of six membrane-spanning α-helices labeled S1 through S6. S5 and S6 were

known to line the pore, and S4 was known to form the largest part of the voltage-

sensing apparatus because of its five arginines and two lysines which carry a positive

charge at physiological pH [9].

3Hydropathy plots are tools to analyze the periodicity in the occurrence of hydrophobic aminoacids in a protein. They can be used to predict secondary structures of a protein as well as whichstretch of a transmembrane protein is sequestered in the hydrophobic membrane.

5

Figure 1.3: KcsA crystal structure. Its four subunits are colored differently to make themeasily distinguishable. The α-helical regions of the channel are drawn as cylinders. Thebackbone atoms of the selectivity filter are also drawn, and the oxygen atoms, all facingthe pore, are colored red. Left Top-down view looking into the pore from outside the cell.Right Side view from within the plane of the membrane.

In 1998, the crystal structure of a bacterial ion channel called KcsA was published

[10]. I have included the structure in Figure 1.3. KcsA is a much simpler channel than

Shaker because it lacks the voltage sensing apparatus of S1–S4. However, KcsA shares

a very high sequence homology with Shaker’s S5, S6, and pore regions, and as such

provided insight into the detailed workings of the selectivity filter and the pore. The

selectivity filters of all potassium channels share a motifs of GYG (glycine-tyrosine-

glycine). The crystal structure showed that the filter is formed by the carbonyl

oxygens of the protein’s backbone rather than the amino acid side chains, and the

mechanism of its remarkable specificity was soon elaborated [6] & [11]. The pore lining

region of the schematic in Figure 1.2, is adapted from the KcsA crystal structure.

Connected to the transmembrane domain discussed above is an intracellular do-

main called the “tetramerization domain” because it lends stability to the tetramer

and specificity for tetramerization within each family of potassium channels [12]. At

6

Figure 1.4: KvAP crystal structure. The views are analogous to Figure 1.3. In addition,the residue corresponding to A359 in Shaker is drawn in orange. A359 is located at junctionbetween S4 and the S3–S4 linker in Shaker. The measurements in Chapter 4 use a dyeattached to this site.

the end of the N-terminal domain is the “inactivation ball” whose function is to lit-

erally plug the entrance to the channel roughly 10–100 ms after it opens [13]. The

inactivation ball is important to the channel’s role in generating the action potential

of neurons because it helps to limit the duration of the action potential by halting the

flow of ions across the membrane. The orange box in Figure 1.2, left demarcates the

region of the protein involved in transducing the conformational changes of gating,

inactivation, and interacting with other proteins.

In the next section I will turn from the broad features of Shaker to focus on the

voltage sensor, since interest in it motivates the work presented in Chapters 3 and 4.

1.2.2 The voltage sensor

In order for Shaker to function as a voltage gated channel, part of the channel must

detect the electrostatic potential across the membrane and move when it changes.

The motion of the voltage sensor is coupled to the “gate” which opens to permit the

flow of ions. Because of the large number of charges on S4, it is believed to be the

voltage sensor, and the intracellular ends of S5 and S6 function as the gate.

When the voltage sensors of all four subunits move, they bring approximately 12–

7

13 elemental charges, i.e. electron charges, across the membrane [14]. The motion

of the voltage sensor is therefore detectable as a transient current which rises quickly

and falls again in ∼10 ms, called the “gating current,” as it is a consequence of thechannel’s gating. The gating current is 200× smaller than the current from the flowof potassium ions through the pore, so the ionic current must be blocked in order to

see it. We use a variant of Shaker with a mutation in the pore, W434F4, which blocks

the ionic current.

The gating current can be used to monitor the opening and closing of the ensemble

of channels in the membrane. Shaker channels are almost completely closed in when

the interior of the cell is 90mV below the exterior of the cell and nearly all open when

the interior is at the same potential as the exterior.

The traditional model for the motion of the voltage sensor is a large translation

across the cell membrane [15]. Using lanthanide-based resonance energy transfer, a

fluorescence technique for measuring distances, we were able to measure the distance

between homologous sites on the four subunits of Shaker. We found no evidence for

a large translation. Rather it appears that the helices of S1–S4 are tightly-packed,

and S4 undergoes a rotation to transport its charges from one side of the membrane

to the other [16].

Figure 1.2 depicts a model which grew out of the work presented in Chapter 3.

In the model, S1–S3 contact the membrane and shield S4 from it. S4 is exposed to

the solution on either side of the cell by invaginations, or vestibules, which extend

into the protein itself. Notice that S4 spans the membrane but is both isolated from

it by the surrounding protein and exposed to the aqueous solution on both sides of

the membrane by invaginations in the protein surface. Hence, the field across the

membrane becomes a field across S4, and S4 is able to move charge across the field

by rotating from a position exposing it to one vestibule to a position exposing it to

4The notation for point mutations is explained in Appendix A.

8

the other, and no large translation is needed.

Interest in the voltage sensor has redoubled this year because of the publication of

the X-ray crystal structure of a voltage-gated potassium channel from archaebacteria,

KvAP [17] & [18]. Figure 1.4 depicts the crystal structure for easy comparison with

Figure 1.3. The sequence of KvAP is similar to Shaker in the voltage-sensing region,

with the notable exception that it lacks the sequence between the S3–S4 linker which is

where we found the most interesting distance measurements discussed in Chapter 3.

For reference, I have colored a residue homologous with A359 in Shaker which we

study extensively in Chapter 4 in orange. A359 is at the top of S4. In Figure 1.4 it

appears at the tip of a “voltage sensing paddle” which is proposed to undergo a large

translation across the membrane during channel activation.

The crystal structure has been met with some puzzlement by the community of

ion channel researchers [19]. The location of the “paddles” exposes the charges on

S4 to the membrane, a configuration which was thought to be unlikely before the

crystal structure because the energetic cost of moving charges from a medium of high

dielectric constant, ²water = 80, to a medium of low dielectric constant, ²lipid = 2,

was thought to be prohibitive. It is believed that the antibodies raised to the voltage

sensor which enabled the protein to be crystallized also disrupted its structure. More-

over, the model proposed for the motion of the paddles involves a large translation,

contradicting our data. Thus, despite the crystal structure, there is still room for in

situ structural studies of voltage gated K+ channels.

1.3 Single molecule spectroscopy and myosin V

After working with the Shaker potassium channel, I redirected my efforts to studying

myosin V at the level of single molecules. In recent years, the ability to study single

fluorescent molecules has become very popular [20]. Single molecule experiments can

reveal distributions in the behavior of biological molecules which are averaged out in

9

an ensemble measurement.

Single molecule experiments require not only modern optics and detectors to cap-

ture as many photons as possible from a single fluorophore, but also high-quality

fluorophores to emit as many photons as possible. A good fluorophore will have a

large extinction coefficient to absorb excitation photons efficiently, a high quantum

yield to re-emit the absorbed energy as photons rather than as heat. In the next

section I introduce a fluorophore which is not among the best by these measures, but

which possesses other significant advantages over conventional dyes which make it

interesting and useful.

1.3.1 The green fluorescent protein

In this section I introduce the green fluorescent

Figure 1.5: Crystal structureof eGFP. The peptide backbone isgreen and the chromophore is yel-low. The structure approximatesa cylinder 42 Å long and 24 Å indiameter.

protein (GFP). It is an important enough protein

in this thesis, where it appears in more than one

context as a fluorescent probe, to merit its own

introduction, even though it has not been an object

of study in itself. I will discuss some of the major

advantages it has as a fluorescent probe in biology

as well as some of its complementary shortcomings.

I begin with its historical origins.

GFP is found natively in the jellyfish, Aequorea,

where it acts as a companion molecule to aequorin,

a chemiluminescent protein which emits in the blue,

∼ 470nm, and excites GFP fluorescence via energytransfer (see Chapter 2) so that the jellyfish glows green, with a peak at 507 nm.

Fundamental questions remain about what evolutionary advantage glowing green has

over blue, why a system of two proteins involved to produce green light instead of

10

evolving a green aequorin, and why the jellyfish should want to glow in the first place

[21].

In 1962, GFP was isolated and shown to be distinct from aequorin [22], but it

wasn’t until 1992 that its gene was cloned [23]. In another two years, the gene was

expressed in non-native systems [24] and [25] demonstrating that the chromophore

forms spontaneously and does not require specific enzymes for its formation. Another

two years later, GFP’s photophysical properties were improved by the discovery of

mutations F64L and S65T [26] which enhance the absorption at 475 nm at the ex-

pense of the absorption at 395 nm, making it more suited for use with the 488 nm

line from the common Ar-ion laser, as well as enhancing the overall brightness of

the fluorescence. The mutant was called GFPmut1 by its discoverers, but today is

commonly referred to as “enhanced GFP,” or eGFP.

Since then, interest in using GFP as a fluorescent marker for studies on cells as well

as in vitro has spread rapidly throughout the biological community. This interest has

been enhanced by the discoveries of many mutations which alter the color or tailor

other photophysical properties to suit experimental needs. In applications where

brightness and photostability are paramount, such as single-molecule work, eGFP

remains a top choice.

Four advantages to labeling with GFP are given in [27], to which I add a fifth:

1. Some proteins are easily inactivated by chemical modification, i.e. attaching an

exogenous dye.

2. Chemical labeling requires the availability of milligram quantities of highly pu-

rified protein, and often leads to large losses.

3. Chemical labeling gives a statistical mixture of proteins with differing numbers

of fluorophores, complicating data analysis.

4. The GFP methodology is easier to use than labeling with exogenous dyes. Pro-

11

teins may be tested immediately after column purification.

5. The mutagenesis required to remove endogenous cysteines and insert one at an

interesting position (cf. Section 2.1) can kill its activity.

Despite these advantages, GFP’s most significant drawbacks relative to conven-

tional fluorophores are its photostability and brightness. It has a high quantum yield

(0.6), but its molar extinction coefficient is only 60,000 M−1cm−1 [21]. And whereas

Cy3 emits a million detectable photons before bleaching [28], a long-lived GFP will

only emit ∼100,000. Since single molecule work requires the ability to collect a largenumber of photons from a molecule, the suitability of eGFP to single molecule work

has heretofore been limited.

GFP has been used to study the molecular motor kinesin at the single-molecule

level [29]. However the localization accuracy attained was 40 nm. We have achieved

an improvement of more than four-fold in spatial precision, with time resolution of

0.5 sec, and we have applied our technique to the study of another molecular motor,

myosin V, to which I turn in the next section.

1.3.2 Myosin V

The myosin family of motor proteins is large and diverse, containing fifteen major

classes, and much variation within each class [30]. Myosins are molecular motors

which hydrolyze ATP5 and bind with polymers of another protein, actin, in a cycle

which produces motion of the myosin relative to actin. Figure 1.6 illustrates the this

cycle for the myosin I have studied, myosin V.

Myosin V is called an “unconventional” myosin to distinguish it from “conven-

tional” myosin, myosin II. Myosin V differs from myosin II in its basic function of

being able to walk along actin fibers without dissociating. Myosin II is “conventional”

5Adenosine triphosphate, the “energy currency” of all cells.

12

Figure 1.6: The catalytic cycle of myosin V. Figure taken from [31].

because its abundance caused it to be the first myosin to be studied.6 Myosins are

characterized by a globular motor domain which hydrolyzes ATP and binds to actin,

a neck region which binds shorter peptides, called light chains or calmodulin, and a

tail domain which tethers the molecule to its cargo and positions the head to interact

with actin [30]. In the myosin II family, the shorter peptides have regulatory duties.

In myosin V, the neck region is very long and binds six calmodulins which lend it

structural rigidity. The calmodulin-binding region is characterized by a repeat of

IQXXXRGXXXR [32], where X can be any amino acid. The neck region of myosin V

is three times longer than the neck of myosin II. Figure 1.7, left is a crystal structure

of myosin II. There is no crystal structure of myosin V, but myosin V shares a 40%

sequence homology with myosin II in the head region [33].

Myosin V is a processive motor involved in transporting organelles in the cell [35].

6Myosin II and actin are the most abundant proteins in muscle cells because their interactiongenerates muscle contraction.

13

Figure 1.7: Left The crystal structure of myosin II subfragment 1. The crystal structurecontains one head of the myosin dimer. The head and neck regions are green, and the twoshort peptides which bind to the myosin II neck, the regulatory and essential light-chains,are blue. The N-terminus is highlighted in orange. The cleft near the end of the head is theentry to the catalytic domain. The actin binding domain is located at the end of the headfarthest from the neck. Right Electron micrographs of myosin V bound to actin suggestingthe telemark skier stance [34].

Defects in myosin V lead failures in cargo trafficking in cells which cause neurological

disorders and lightened coat color in mice [33]. Myosin V consists of two identical

heavy chains, each of which form a globular “head” at the N-terminus which contains

an actin-binding domain and a catalytic domain which hydrolyzes ATP. The head

is often referred to as the “motor domain.” The C-terminus forms a ∼24 nm longα-helical domain which binds six calmodulin molecules and probably serves as a lever

arm to amplify small conformational changes in the globular domain and convert

them into processive motion [35, 36, 37, 38]. The two heads are joined at the end of

the calmodulin-binding domain to form a coiled-coil stalk which attaches the motor

domain to its cargo.

Because of the length of the neck region, myosin V can take very large steps of

35–40 nm for each ATP hydrolyzed [38, 39, 40, 41, 42]. Until recently, however, it

had not been conclusively shown what type of motion the heads undergo: whether

14

Figure 1.8: Schematic of hand-over-hand model of myosin V procession when the leadingmyosin V lever arm is straight (left) or has a bend (right). The step size of a dye (greenor purple dots) alternates between small and large: [37 - (x1 + x2)], followed by [37 + (x1+ x2)], where x1 and x2 are the distances from the midpoint between the globular domainsalong the direction of motion, before and after the step, respectively. With a straightlever arm, x1 = x2 ≡ x, and the step sizes are larger and smaller than 0 nm and 74 nm,respectively, (2x′ < 37 nm) for a dye anywhere on the lever arm. In the bent-lever-armmodel, the same stepping pattern is seen for a dye above the bend; however, a dye belowthe bend (purple dot), takes alternating 0nm and 74nm steps (x′1 + x′2 = 37nm). In eithermodel, the catalytic domain step size, as measured by a GFP attached to the N-terminus,is 74 nm.

one head always leads the other in an inchworm fashion, or whether they alternate

positions like a person walking. Recent results from our lab and our collaborators

have shown that though the center-of-mass of the myosin molecule takes 37nm steps,

each head of the myosin takes alternating steps of 0 nm and 74 nm, which can only

be possible if myosin walks hand over hand as in Figure 1.8 [28] & [38].

How the two heads are coordinated in the hand-over-hand mechanism remains an

unresolved question. Nucleotide hydrolysis, actin binding, and lever arm position are

15

likely to be tightly coupled [43]. Myosin V’s unidirectional motion implies there must

be a functional asymmetry between the leading and trailing heads, i.e. the trailing

head must know to step forward and the leading head must know to stay put. One

model of the asymmetry postulates that the rear head is released from actin when

ATP binds and is then thrown forward by strain created previously in the myosin

[31] & [40]. Strain-induced inhibition of ATP-binding to the leading head has been

postulated for kinesin, a different family of two-headed processive motors [44].

The location of this strain is uncertain. An attractive model is the “telemark

skier,” where the front head is bent or kinked, (Figure 1.8, right) [31, 34, 45]. When

the rear head binds ATP and is released from actin, the strain released from the bend

would tend to throw the rear head forward. A sharp bend in the front head was

seen in some negatively-stained electron micrographs of myosin V, where the bend

was interpreted to be just before the lever arm—most likely in the pliant region [46]

between a part of the catalytic domain known as the converter domain and the first

IQ motif of the lever arm [34]. More gentle distortions in the lever arm itself were also

observed, and other myosin V’s appeared to have straight lever arms. The subset of

lever arms that fell into a relatively straight configuration were analyzed in detail in a

later paper which showed that the rear head was in a post power stroke conformation

and the leading head was in a pre-power stroke conformation [45].

An alternative telemark skier model places the bend directly in the lever arm.

This model makes two testable predictions. First, as noted by Forkey, et al. [38], a

dye rigidly attached to a calmodulin below the kink will not reorient during a step,

while a dye placed above the kink will reorient. Figure 1.9 illustrates this point. In

their polarization-sensitive single molecule motility study of myosin V, Forkey, et al.

observed two populations of dyes, one that rotated during a step and one that did

not, although the locations of the dyes corresponding to these two populations were

not determined. Second, the step size of a dye below the kink will alternate between

16

Figure 1.9: In the telemark skier model, the polarization of a dye attached above a kinkwill rotate, but the polarization of a dye below the kink will not. The red arrows representthe transition dipole of a dye attached to the neck region of myosin V. The blue arrowindicates that the trailing head has become the leading head, and a step has been taken.

74 nm and 0 nm—the same as a dye on the globular domain—while the step size of

a dye above the kink will alternate between a large step that is less than 74 nm and

a small step that is more than 0 nm as in the right side of Figure 1.8. In contrast,

the telemark skier configuration in which the bend is in the converter domain below

the lever arm, as Walker, et al. have suggested [34], leads to steps of < 74 nm and

> 0nm for a dye anywhere in the lever arm—the stepping pattern characteristic of a

straight lever arm.

Yildiz, et al. have developed a technique called FIONA, for Fluorescence Imaging

with One Nanometer Accuracy, with which the position of a dye could be located

with 1.5 nm precision and sub-second temporal resolution [28]. Using FIONA, they

showed that a dye on a calmodulin bound to the neck region of myosin V could take

steps of 0 and 74nm. Assuming the two heads also take 74nm steps, this result favors

17

the second telemark skier model in which the lever arm is bent. However, because the

method of labeling employed by Yildiz, et al. did not allow them to place a dye on

the globular domain, a direct measurement of the globular domain step size remains

necessary to confirm the model.

We have extended FIONA to enable nanometer localization of single eGFPs and

to measure the step size of the globular domain of myosin V. Others have previously

imaged single GFPs bound to kinesin [47], and 30nm localization with 5msec temporal

resolution has been achieved [48]. Here, we have constructed a heterodimeric myosin V

which has an eGFP linked to the N-terminus of one of the two globular domains and

with it have achieved 4–10 nm localization of single eGFP-myosins with sub-second

temporal resolution and observation times of typically ∼25 sec and occasionally aslong as 75 sec. Control experiments using a nanometric stage verify our ability to

accurately measure step sizes. ATP-induced myosin V steps are also measured. We

find the globular domain takes 74 nm steps, which, when combined with the results

of Yildiz, et al. supports a telemark skier model with a bend in the lever arm of the

leading head.

18

Chapter 2

Applications of luminescentlanthanide chelates

This chapter provides technical background for the one that follows and culminates

with experimental results from the beginning of my graduate career. In Chapter 3,

I present results obtained through Lanthanide-based Resonance Energy Transfer, or

LRET, which uses luminescent lanthanide chelates developed by my advisor [49] & [50]

to measure distances on the length scale of proteins. The physical theory needed to

apply LRET was first worked out for conventional fluorophores, and is called Fluores-

cence Resonance Energy Transfer, FRET. In order to understand LRET’s advantages

over FRET, I begin by explaining the theory of FRET, followed by a description of

the advantages of LRET important to the results presented in Chapter 3, and use

these to motivate the experiments presented in the last section of this chapter.

2.1 Fluorescence resonance energy transfer

(FRET)

FRET is a technique for measuring distances on the scale of a protein, ∼2–10 nm.Its great advantage is that it enables distances shorter than optical wavelengths,

450 nm ≤ λ ≤ 650 nm, to be measured with visible light. FRET is done by attachingtwo different fluorescent dyes to the protein under study. One dye is called the “donor”

because it is excited by the experimenter and transfers, or donates, its energy to the

19

NH ONH

ON

CO2-

N

N

CO2-

CO2- O

R

Tb3+

NOO

R’

S

cys

N

R’

OO

S-

cysH+

Figure 2.1: Left The structure of the lanthanide chelate. Eu3+-DTPA-cs124 has the samestructure with the Tb3+ replaced with Eu3+. R designates another organic group, such asa maleimide or a protein. Right The structure of the maleimide group before and after thenucleophilic attack of the sulfur on a cysteine side chain. R′ represents the fluorescent dyebeing attached to the protein.

other dye, called the “acceptor,” with an efficiency that depends on the distance

between them. The distance is calculated by measuring the energy transfer efficiency,

as described below.

The method of attaching the dyes can vary. In the experiments of Chapters 3 and 4,

fluorescent dyes are attached to the Shaker potassium channel by using site-directed

mutagenesis to insert a cysteine residue into the protein at the site of interest and

then attaching a dye with a chemical group which reacts to the sulfhydryl group at

the end of the cysteine. Figure 2.1 illustrates the chemical reaction which occurs.

At pH > 6, the sulfur at the end of the side chain of the cysteine is de-protonated

and can perform a nucleophilic attack on the double bond of the maleimide, form-

ing an irreversible covalent attachment between the protein and the fluorescent dye.

The method of directing the two dyes to different, but specific cysteines on the pro-

tein varies for each protein under study. The experiments described in the next two

chapters avoided this difficulty by exploiting the symmetry of the channel.

The theory for FRET was derived by Förster [51], but the treatment I present

here is adapted from [52]. Energy transfer occurs via the interaction of the electric

dipole moments of the donor and acceptor. The rate at which energy transfer occurs,

ket is given by Fermi’s Golden Rule:

20

ket ∝∣∣∣∣∣〈D

∗, A|µD ·µAR3

− 3(µD ·R)(µA ·R)R5

|D,A∗〉∣∣∣∣∣2

(2.1)

where D and A refer to the donor and acceptor, µD and µA are their electric transition

dipole moments, R is the distance vector between the two dyes, and ∗ denotes the

excited state.

The efficiency of energy transfer,E, is given by

E =ket

ket + knd

=1

1 + knd/ket(2.2)

where knd is the rate at which the donor de-excites through all non-distance dependent

pathways, such as internal vibrations. Equation 2.1 allows us to get a distance from

Equation 2.2. Combining all of the physical constants from knd and Equation 2.1 into

one quantity, Ro, we are able to write the energy transfer efficiency in the common

and useful form,

E =1

1 + (R/Ro)6 . (2.3)

The sixth-power dependence on R comes from squaring the dipole-dipole energy in

Fermi’s Golden Rule. Because E falls so steeply with R, it is most sensitive for

distances near Ro. Ro depends on the identity of dyes chosen for the donor/acceptor

pair through

Ro = (8.79× 10−5JqDn−4κ2)1/6 (in Å), (2.4)

where qD is the quantum yield of the donor, n is the index of refraction of the medium

separating the two dyes, κ2 is a factor which depends on the orientation between the

dipoles of the two dyes, and

21

J =

∫²A(λ)fD(λ)λ

4dλ∫

fD(λ)dλ. (2.5)

J is known as the normalized spectral overlap integral because it depends on the

overlap of the molar absorption spectrum of the acceptor, ²A(λ), and the fluorescence

spectrum of the donor, fD(λ).

All of the quantities in Ro are usually well-known when a FRET experiment is

devised. J may be determined quite easily from spectra which are similarly easy

to obtain. In fact, the steep dependence of E on Ro encourages the investigator to

consider J when choosing a donor-acceptor pair. The quantum yield of the donor,

qD, can be measured or approximated, and n is usually taken to be n = 1.33, the

index of refraction of water.

Contact with measurable quantities is made through the relation,

E =(1− τDA

τD

), (2.6)

where τDA is the excited state lifetime of the donor in the presence of the acceptor, i.e.

on a doubly labeled sample, and τD is the lifetime of the donor. Since the acceptor

always shortens the lifetime of the donor by providing an additional de-excitation

pathway, E < 1. Combining Equation 2.6 with Equation 2.3, we get

R = Ro

(τDτDA

− 1)−1/6

. (2.7)

The greatest source of error in a FRET experiment is κ2 in Equation 2.4, which

comes from the dot products in Equation 2.1. κ2 can range from zero when the

transition dipoles of the donor and acceptor are perpendicular to four when they are

collinear. κ2 is usually approximated as 2/3, which is its value when the dyes are

free to completely re-orient during the excited state lifetime of the donor. Since the

donor and acceptor are attached to a protein, they are often not completely free to re-

orient, and an exact measurement of κ2 can be difficult or impossible, resulting in an

22

error in the computed distance. If the dyes can partially reorient during the donor’s

excited-state lifetime, then limits can be placed on the range of possible values of κ2.

However, the uncertainty in R caused by uncertainty in κ2 can be greatly reduced by

using a lanthanide atom as the donor, a technique we turn to now.

2.2 Lanthanide-based resonance energy transfer

(LRET)

LRET is a variation of FRET in which the donor is a lanthanide atom instead of a

conventional fluorophore. The acceptor in LRET is a conventional acceptor, usually

an organic fluorophore. Förster theory also applies to LRET with qD taken as the

quantum yield of the lanthanide atom.

In Chapter 3, we measure the lifetime of the “sensitized” emission of the acceptor,

τAD. Sensitized emission consists of the photons emitted by the acceptor after being

excited by energy transfer from the donor. Because the excited state lifetime of a

lanthanide chelate (1.55 ms for Tb3+-DTPA-cs124) is six orders of magnitude longer

than the lifetime of a conventional organic dye, the decay in sensitized emission follows

the same time course as the decay in donor luminescence in an LRET experiment,

and τAD is equal to τDA in Equation 2.6. Energy transfer measured from sensitized

emission using LRET has several advantages over conventional FRET [52]:

• spectral discrimination against background In a conventional FRET ex-periment, the emission spectra of the donor and acceptor often overlap, leading

to contamination from donor fluorescence of the signal from sensitized emission.

In contrast, since the donor in an LRET experiment is an atom rather than a

molecule, its emission spectrum is nearly discrete.1 The acceptor can be chosen

to emit in a range of wavelengths where the donor is dark, thereby eliminating

1The peaks are typically 10 nm wide, significantly narrower than emission from organic fluo-rophores, although larger than emission from an isolated atom because the electric field from theDTPA causes broadening via the Stark effect.

23

contributions of donor luminescence to the measured sensitized emission.

• temporal discrimination against background When measuring E by sen-sitized emission, the major source of background is “prompt” fluorescence from

acceptors not excited by energy transferred from a donor. Two sources of

prompt fluorescence are acceptors with a paired donor that are excited by the

light used to excite the donor and acceptors without a paired donor. Direct ex-

citation of the acceptor is unavoidable in most energy transfer experiments, and

labeling with multiple acceptors is a consequence of the labeling protocol used

in the next chapter. In conventional FRET, the donor and acceptor usually

have excited state lifetimes of the same order of magnitude, making temporal

discrimination against prompt fluorescence difficult. In LRET however, be-

cause the lanthanide lifetime is so much longer than the acceptor lifetime, the

prompt fluorescence has finished before the donor begins to transfer its energy,

and time-gated detection can be used to eliminate prompt fluorescence, leaving

fluorescence due to energy transfer with very low background.

• single-exponential lifetimes Lanthanide chelate donor lifetimes can be single-exponential, permitting the resolution of heterogeneous populations of donor-

acceptor pairs. Many conventional fluorophores do not have single-exponential

decays. The labeling scheme used in Chapter 3 necessarily produces a hetero-

geneous population of donor-acceptor pairs.

• orientation factor The final advantage of LRET over FRET important tothe next chapter is the reduction in uncertainty from κ2. When the donor

emission is intrinsically unpolarized, 1/3 < κ2 < 4/3, and even if the acceptor

is completely immobile, the uncertainty in E from κ2 is reduced to at most 11%.

The uncertainty will be even lower if the acceptor has some rotational freedom.

The structure of the lanthanide chelate is shown in Figure 2.1. It consists of three

24

parts, the “antenna,” the chelator, and the lanthanide. The antenna is carbostyril-

124, or cs124. It consists of two conjugated aromatic rings. It has a maximal ab-

sorption at 328nm. Its purpose is to increase the effective absorption cross-section of

the lanthanide by four orders of magnitude from < 1 M−1cm−1 to 12, 000 M−1cm−1

[50] by absorbing excitation photons and transferring their energy non-radiatively

to the lanthanide. The chelator is diethylenetriaminepentaacetic acid, or DTPA. Its

purpose is to hold the lanthanide in close proximity to the antenna and to shelter it

from water, whose OH bonds quench the lanthanide’s emission [53]. The “R” at the

end of the DTPA refers to an organic molecule, either a chemically reactive group for

attaching the molecule to a protein, or else the protein itself, once it’s attached.

That lanthanide emission is unpolarized was assumed for many years because

the luminescent transitions were believed to occur between highly degenerate, high

angular momentum states [54], but to put LRET experiments on solid footing the

polarization of lanthanide emission needed to be measured directly. The next section

describes measurements of the anisotropy of two lanthanide chelates.

2.3 Anisotropy of lanthanide chelates

In order to claim that the uncertainty in κ2 in our LRET measurements is low, we

needed to measure the anisotropy of the lanthanide chelates to show that it is nearly

zero. The anisotropy is a measure of the extent to which the emission of a fluorophore

is polarized [2]. It is defined as:

r =I‖ − I⊥I‖ + 2I⊥

, (2.8)

where I‖ and I⊥ are the intensities of the fluorescent light with polarizations parallel

and perpendicular to the polarization of the excitation light. The anisotropy may best

be understood with reference to Figure 2.3. The excitation light must be polarized

perpendicular to the plane defined by the excitation and observation pathways, i.e.

25

Figure 2.2: Left Tb3+-DTPA-cs124 anisotropy in red, spectrum in green. Right Eu3+-DTPA-cs124 anisotropy in red, spectrum in green.

perpendicular to the x-y plane, along the z-axis. Notice that if the excitation light

were polarized parallel to the optics table, the two emission polarizations would both

be perpendicular to the excitation polarization, and hence their intensities must be

the same. This observation is useful to correct for any polarization bias in the de-

tecting apparatus, and it illustrates the importance of the geometry of an apparatus

to measure the anisotropy.

Many conventional fluorophores have linearly

Figure 2.3: Geometry of ananisotropy measurement.

polarized emission because of the dipole nature of

the fluorescent transition. However, even a conven-

tional fluorophore will have a very low anisotropy

in aqueous solution because between the time it ab-

sorbs a photon and emits one of lower energy, it will

have diffused rotationally so that the polarization

of the emitted photon will have no relation to the

polarization of the one absorbed. This effect is even more pronounced for lanthanides

because their excited state lifetimes are six orders of magnitude larger than those of

conventional fluorophores. When two dyes are used for FRET, however, their mo-

bility is restricted by their attachment to the protein under study, which becomes a

26

source of error as discussed in the preceeding section.

To measure the inherent anisotropy, i.e. the anisotropy of an immobilized lan-

thanide chelate, samples of ∼ 1 µM Tb3+-DTPA-cs124 and Eu3+-DTPA-cs124 wereprepared in >90% glycerol. Glycerol forms a glass at temperatures below −80 ◦C. Toachieve such low temperatures, samples were loaded into quartz NMR tubes (Wilmad

Glass) and mounted in a custom-built liquid nitrogen dewar with a quartz bottom.

It was necessary to use quartz instead of glass because glass will not transmit the

337 nm light from the nitrogen laser used to excite the cs124. The dewar permitted

experiments to be performed with the sample immersed in liquid nitrogen, well below

the glass transition temperature of glycerol.

The spectrometer consisted of a pulsed nitrogen laser (Laser Photonics) with a

dye laser attachment, a spectrometer (Triax 320M, SPEX), with a liquid nitrogen

cooled CCD. A Glan-Thompson polarizer was used to ensure the polarization purity

of the excitation light, and polymer sheet polarizer was used in the emission path

to select I‖ or I⊥ for detection. This setup permits the detection of I‖ or I⊥ for the

entire emission spectrum of the dye at once, enabling the calculation of the anisotropy

as a function of wavelength. To improve the signal-to-noise ratio, the spectrum was

integrated for many laser pulses, and to compensate for photobleaching of the dye,

spectra were acquired in the pattern of I‖, I⊥, I⊥, I‖, and then added appropriately.

The anisotropy of the two lanthanide chelates as a function of wavelength is shown

in Figure 2.2. Tb3+-DTPA-cs124 is unpolarized, as we expected, but Eu3+-DTPA-

cs124, surprisingly, has a strongly wavelength-dependent anisotropy. The emission

of Tb3+-DTPA-cs124 could be unpolarized because the process of transferring en-

ergy from the cs124 antenna to the Tb3+ loses information about the polarization of

the excitation photons, or it could be unpolarized because the emission of Tb3+ is

inherently unpolarized.

To distinguish between these two possibilities, we excited the ∼ 490nm transition

27

Figure 2.4: Tb-DTPA anisotropy in red, spectrum in green.

of Tb3+-DTPA directly with the dye laser attachment to the nitrogen laser. The

chelator, DTPA, was retained to increase the brightness of Tb3+ by expelling OH

groups from the solvent and to surround the lanthanide with the same environment

that it has in the full chelate with the antenna. The laser was tuned to maximize

luminescence from the lanthanide.

Figure 2.4 shows that the anisotropy of Tb3+-DTPA is very close to zero. The

anisotropy of the manifold centered at 490nm is difficult to measure because scattered

laser excitation contributes to the measured intensity at those wavelengths and is

polarized. However, the anisotropy of the 490nm manifold, as well as the anisotropy

of Eu3+-DTPA were measured by a younger student in our lab, Jeff Reifenberger [55],

and the 490 nm manifold of Tb-DTPA was found to be unpolarized.

Having shown that the emission of the Tb3+-DTPA-cs124 donor used in the energy

transfer experiments of the next section is inherently unpolarized, and the uncertainty

in κ2 is consequently minimized, we were able report very small (∼ 1 Å) distancechanges as a consequence of conformational changes in the Shaker potassium channel.

28

Chapter 3

The motion of the voltage sensor ofShaker detected by LRET

In this chapter I describe some experiments that were only possible because of the

advantages of LRET discussed in the previous chapter. We use LRET to obtain the

first measurements of distances between specific homologous sites in the S3–S4 region

on different subunits of Shaker in situ, and find that changes in those distances are

correlated with the motion of the voltage sensor detected by electrophysiology. We

find that the voltage sensor does not undergo a large conformational change such as a

translation perpendicular to the plane of the membrane. Rather, the results presented

here suggest an exciting new model for the conformational change in gating, namely

that the voltage sensor undergoes a rotation. These results have been published in

Nature [16].

3.1 Materials and methods

3.1.1 Sample preparation

Protocols for preparing Shaker channels for study are given in Appendix B. Briefly,

Shaker channels are prepared by injecting mRNA for the channel into oocytes1 har-

vested from Xenopus laevis frogs. This procedure accomplishes two things: the oocyte

manufactures the channel and provides a membrane in which to study it. It has the

1Oocytes are the female gametes, or egg cells.

29

P

N

C

+

+

+

+

+

--

-

S1 S2 S3 S4 S5 S6+

+

270

273

346 351

363

353 425352

RRSSCC

SSRR AASSRR AARR AA

Terbium−−maleimide

Fluorescein−−maleimide

Figure 3.1: Left Schematic of a single subunit of Shaker showing the locations of themutations studied as well as the native charged residues. Right Schematic of the twodistances measured by LRET using stoichiometric labeling. The most common distributionof donors and acceptors is shown.

advantage that oocytes are macroscopic, ∼ 1 mm in diameter. Oocyte are thus easyto manipulate and contain many Shaker channels, leading to robust signals. Indeed,

oocytes are the standard cell used in electrophysiological studies of many ion channels.

3.1.2 Labeling with fluorescent dyes

Figure 3.1 illustrates the locations of the mutations made to attach fluorescent dyes.

The figure on the left is a schematic of one subunit of the channel showing the six

transmembrane segments deduced from hydropathy analysis and accessibility studies.

The figure on the right illustrates that for each cysteine mutation, two distances will

be measured: one between adjacent subunits (RSC) and one across the pore (RSA).

Moreover, because of the tetrameric symmetry of the channel, the dyes lie near the

corners of a square and the two distances are expected to be related by√

2.

Channels were labeled by exposing oocytes to a 4:1 mixture of Tb3+-DTPA-cs124-

maleimide and fluorescein-maleimide. The distribution of donors and acceptors is

therefore determined by the stoichiometry of the labeling mixture. Assuming that

both dyes were equally reactive and that all channels were labeled completely, then

the labeling frequency is obtained from the binomial distribution. 41% will have four

30

Figure 3.2: Apparatus to combine two-electrode voltage clamping with fluorescence.

donors, 41% will have three donors and one acceptor, 15% will have two donors and

two acceptors, 2.6% will have three acceptors and one donor, and 0.16% will have

four acceptors.

3.1.3 Two electrode voltage clamp

There are many techniques for controlling the electrostatic potential across a cell

membrane. All of them are referred to as “voltage clamping” because the instrument

holds, or “clamps,” the voltage at a value chosen by the experimenter by injecting the

necessary current to maintain the voltage. The “patch clamp,” for which the 1991

Nobel Prize in Physiology and Medicine was awarded, is perhaps the most famous

voltage clamp technique. We primarily use a simpler technique, the two-electrode

voltage clamp (TEV).2

In addition to allowing us to control the potential, the voltage clamp enables us

to measure the ionic currents of potassium flowing through the channel and gating

currents when we block the ionic current. (See Chapter 1.2.2.) The voltage clamp

2The experiments discussed in this section were actually performed using the “cut open oocyte”voltage clamp, but the two-electrode clamp is discussed for simplicity and because the distinctiondoes not affect our results.

31

enables us to monitor the health of a cell (since cells with leaky membranes cannot

be clamped), to detect the presence and activity of the channels of interest electri-

cally, and to correlate our fluorescence measurements with the state (open or closed)

of the channel. The voltage clamp (CA-1B, Dagan) is set up on an inverted flu-

orescence microscope (IX-70, Olympus) for simultaneous fluorescence and electrical

measurements.

The name TEV refers to the two electrodes (I and V in Figure 3.2) needed to

control the potential inside the oocyte, one to monitor the membrane potential, and

the other to inject current needed to clamp the membrane at the desired potential. A

pair of agar bridges (P1 and P2) containing a high-salt solution monitor the potential

of the solution surrounding the oocyte and inject current to clamp the membrane at

the desired potential. The interior of the cell is held at “virtual ground,” while the

potential of the solution bathing it is changed.

3.1.4 Fluorescence detection

As indicated in Figure 3.2, experiments were performed on an inverted microscope

so that the microscope objective is underneath the oocyte. Excitation light from the

pulsed nitrogen laser used to excite donor fluorescence was separated from fluores-

cent light from the sample, and bandpass filters were used to select the sensitized

emission of the acceptor. Photons were collected in a time-dependent manner by a

photomultiplier tube to capture the decay of the luminescence.

3.2 Results and discussion

The basic experimental procedure was to hold the membrane at −90mV, a potentialwhere most of the channels are closed, and pulse the membrane to a higher potential

in 10mV increments in order to open the channels. The laser is fired 50ms after the

start of the pulse so that all of the gating charge has stopped moving. The voltage

32

−120 mV: τ = 73 µµss

−90 mV−120 mV

50 mV......

Laser

Acquisition

50 ms

10 ms

50 mV: τ = 144 µµss100 µµss

Figure 3.3: Left The lifetime of the sensitized emission changes visibly with voltage.Right The position of S346C (RSC) closely follows gating charge (Q).

pulse is maintained for another 10ms in order to allow all of the luminescence from the

sample to decay. The left side of Figure 3.3 shows that for the S346C mutation, the

time constant of the luminescence decay changes from 173 µs when the membrane is

pulsed to 50mV to 65µs when the membrane is pulsed to −120mV. When distancesare computed from the lifetimes using Equation 2.7 and compared to the fraction

of gating charge displaced, Q, for each pulse potential, the resulting curves nearly

overlap (Figure 3.3, right). Comparison with Figure 3.1 shows that S346C is in the

S3–S4 linker, quite far from the S4. That the motion of this residue follows the gating

charge motion so closely suggests that the linker must be fairly rigid. Indeed, it was

shown later [56] that the S3–S4 linker region is α-helical.

The distances measured by LRET are summarized in Figure 3.4. RSC and RSA

correspond to two components of the multi-exponential decay of the sensitized emis-

sion. Because the two distances are not independent, but related by the Pythagorean

theorem because of the tetrameric symmetry of the channel, a value R′SA can be cal-

culated from RSC and compared to the measured RSA as an internal control. As the

table shows, the two distances are very similar for every residue, demonstrating the

internal consistency of the data and that LRET can be used to measure absolute

distances.

33

As a control experiment, the mutation F425C was introduced to compare a dis-

tance obtained from LRET on Shaker to the distance between homologous sites in the

crystal structure of KcsA. The distance measured via LRET, 30 Å agrees very well

with the distance measured on the crystal structure, 29 Å. This is a valid comparison

because the pores of both KcsA and Shaker bind toxins which are specific to the

structure of the pore [57], indicating that their pores have similar structures.

Stoichiometric labeling produces an ensemble of channels labeled with a distri-

bution of donors and acceptors. (See Section 3.1.2.) For the reasons described in

Section 2.2, the channels with four donors and four acceptors do not contaminate the

signal. The small subpopulation of channels with two acceptors per donor contributes

an error of ∼ 1.4 Å for a distance of 28 Å, which is nearly eliminated when measuring

0.5

n=6

0.5, 0.6

n=7

0.5, 0.7

n=5

0.4, 1.1

n=3

0.5, 1.6

n=13

0.6, 1.3

n=5

0.5, 0.6

n=9

0.7, 1.1

n=9

standard error

RRSC, RRSA

+5.4%, −−3.3%

+4.5% −−2.7%

+6.0%, −−3.9%

+5.3%, −−3.3%

+5.8%, −−3.7%

+6%, −−3.8%

+5/5%, −−3.4%

anisotropy

constraints

2930F425CPore

454532V363CS4

37−−42

40

41

41

37−−42

41−−42

40

42−−44

26−−30

28

29

29

S346C

S351C

S352C

N353C

S3−−S4

loop

43

46

45

47

31

32

D270C

P273C

S1−−S2

loop

R ŚA(Å)

RRSA(Å)

RRSC(Å)

sitelocation

0.5

n=6

0.5, 0.6

n=7

0.5, 0.7

n=5

0.4, 1.1

n=3

0.5, 1.6

n=13

0.6, 1.3

n=5

0.5, 0.6

n=9

0.7, 1.1

n=9

standard error

RRSC, RRSA

+5.4%, −−3.3%

+4.5% −−2.7%

+6.0%, −−3.9%

+5.3%, −−3.3%

+5.8%, −−3.7%

+6%, −−3.8%

+5/5%, −−3.4%

anisotropy

constraints

2930F425CPore

454532V363CS4

37−−42

40

41

41

37−−42

41−−42

40

42−−44

26−−30

28

29

29

S346C

S351C

S352C

N353C

S3−−S4

loop

43

46

45

47

31

32

D270C

P273C

S1−−S2

loop

R ŚA(Å)

RRSA(Å)

RRSC(Å)

sitelocation

Figure 3.4: Table of distances. The meanings of the various Rs are described in thetext. Where different distances were found in the open and closed states of the channel, thedistance in the closed state is given in bold italics. The number of oocytes used for eachmeasured distance is n. The last column gives limits on the possible error due to κ2 basedon measurements of the anisotropy of the acceptor at each site.

34

RESTING ACTIVE OPEN

RRRR

rraa

−120 −60 6000

R (A)

V (mV)

RRrr

RRaa

RRrr

oo

Figure 3.5: A bell-shaped distance versus voltage curve would result from a large trans-membrane displacement of the voltage sensors.

changes in distance, e.g. as a function of voltage.

The voltage sensors of each subunit are believed to move independently except

for a final cooperative transition [58] immediately preceeding channel opening. If

the motion of the sensors involved a large translation perpendicular to the plane of

the membrane as in Figure 3.5, we would expect to see an increase in the distance

between labels at intermediate potentials where not all of the voltage sensors are in

the activated state followed by a decrease in distance at higher potentials when all

of the voltage sensors have moved. The plot of distance versus voltage would thus

have a bell shape. Such a translation as large as 16 Å has been proposed [59] & [60].

This shape is clearly absent from Figure 3.3. Since the distance between a residue in

S4, V363C, does not change with voltage, we conclude that a large translation of S4

perpendicular to the membrane is unlikely.

Voltage-dependent distance changes were also detected at three adjacent sites near

the top of S4, at S351C, S352C and N353C. The distance changes show a surprising

pattern. The dyes at S351C move away from each other, the dyes at N353C move

35

dist

ance

cha

nge

(Å)

RR50mV

−R−120mV

S351C(n=3)

S352C(n=13)

N353C(n=5)

22

−2

−1

11

00

Figure 3.6: Upper Distance change upon membrane depolarization for three adjacentresidues in the S3–S4 linker. Lower Looking down into the pore so that the tops of the S4sperpendicular to the membrane are visible. A rotation of S4 could produce the distancechanges in the upper figure.

closer together, while the dyes at the residue between them maintain a constant

separation. A simple explanation for this pattern is that the dyes are attached to an

α-helix which rotates as the voltage changes. Figure 3.6 illustrates this model. In

the lower part of the figure, channel opening causes the distance between the black

dots to increase, the distance between the white dots to decrease, and the distance

between the grey dots to stay the same.

36

3.3 The Big Picture

There are two ways in which this work has been supplemented and extended by other

labs since its publication. The first is by in vivo distance measurements akin to ours,

and the second is by the crystallization of a voltage gated potassium channel from

thermophilic bacteria.

When this work appeared in Nature, it was followed immediately in the same

issue by an article from Ehud Isacoff’s laboratory presenting FRET measurements of

distance changes in the S3–S4 region as a function of voltage [61]. As we did, the

Isacoff lab found evidence for a rotation of S4 in distance changes which alternate

in sign between adjacent sites, but oddly enough, their distance changes are in the

opposite direction from ours. Also, the magnitudes of the absolute distances they

measure are larger. They find distances between adjacent subunits to be 50–60 Å,

whereas we find the distances to be ∼ 30 Å. The discrepancy could arise from anumber of sources. Because of the orientation factor, κ2, absolute distances measured

by FRET tend not to be as accurate as absolute distances measured by LRET. Also,

since different dyes were used, their motion relative to the protein may have been

different.

A particular disadvantage of the technique employed by the Isacoff lab is that since

FRET was measured by donor photobleaching, a curve such as the one in Figure 3.3

showing the distance change as a function of voltage is impossible to construct [62].

They were thus unable to reach what is perhaps our strongest conclusion, that there

is little, if any, translational motion perpendicular to the membrane.

Support for our distance measurements came a few months later from Chris

Miller’s group [63] when they published experiments using a clever polymer cross-

linking technique to measure distances from the S4 region to the pore. These mea-

surements agree remarkably well (to within 3 Å) with the the distances we measured.

However, these measurements share the limitation of Isacoff’s of being unable to

37

detect distance changes as a function of voltage.

Earlier this year, a major milestone in the study of voltage gated channels was

reached with the publication of the crystal structure of KvAP, a voltage gated potas-

sium channel from the thermophilic bacterium, Aeropyrum pernix [17]. A picture of

the crystal structure is found in Figure 1.4. KvAP is similar in sequence to Shaker,

but it lacks the S3–S4 linker where the most important residues we studied (S346C,

S351C–N353C) are located. Direct comparisons of the distances we measured with

distances on the crystal structure are therefore imperfect, but they still can be mean-

ingful.

KvAP, the first voltage gated channel to be crystallized, was chosen because it

comes from a thermophilic bacterium with the rationale that it should be more stable

and thus easier to crystallize than most of the channels commonly studied which

come from higher organisms. Also, since the voltage sensor is known to be mobile,

antibodies to it were raised in order to stabilize it, and the channel was crystallized

in the presence of the antibodies. As a result, the crystal structure is acknowledged

by all to be distorted, even by its discoverers. However, opinions are divided on the

extent of the distortion [19].

The crystal structure is inconsistent with all of the distance measurements I have

discussed. In our data, RSA for V363C is 45 Å, but the distance between homolo-

gous residues in the crystal structure (measured using VMD [64]) is 88 Å. Moreover

the model proposed in the crystal structure papers requires a large transmembrane

displacement of S4 [18], whereas our data specifically rule that out.

The most exciting result to emerge from the work presented here is the model of

the rotation of S4. Since the evidence for the rotation is indirect, as it comes from

measurements of linear distances in the S3–S4 linker, we wanted to detect it directly

by placing a probe sensitive to rotation son S4 itself. This desire motivates the work

begun in the next chapter.

38

Chapter 4

Mobility of Shaker in the oocytemembrane

The results of the previous chapter suggest that S4 undergoes a rotation, but they

do not detect the rotation directly. We wanted to do experiments to detect the

rotation directly. However, the slow time scale of gating requires that the entire

channel be rotationally immobile on the time scale of milliseconds lest motion of

the whole protein obscure the movement we wished to detect. Hence, we decided

to do a control measurement to determine whether the channels were mobile before

attempting a rotation measurement.

In this chapter I present experiments using Fluorescence Recovery After Photo-

bleaching (FRAP) which demonstrate the channel to be translationally immobile in

the oocyte membrane and attempt to identify the agent preventing its diffusion. Af-

ter presenting the diffusion measurements, I briefly discuss preliminary results in the

construction of an apparatus to detect the rotation of S4 using a variation of FRAP

sensitive to the orientation of the fluorescent probe known as polarized FRAP, or

pFRAP.

39

4.1 Fluorescence Recovery After Photobleaching

(FRAP)

Fluorescence recovery after photobleaching (FRAP) exploits a property of fluorescent

dyes usually regarded as a liability: their tendency to be destroyed after undergoing a

number of excitation and emission cycles, called photobleaching. In a FRAP experi-

ment, laser light is focused to a spot on the oocyte membrane. The light is modulated

to produce a brief but intense pulse, followed by a dim and much longer probe beam.

The pulse photobleaches most of the dyes in the spot, and the probe excites the

fluorescence of the remaining fluorophores as well as any that diffuse into the spot

to replace the ones which have been destroyed. Usually, many bleach and recovery

cycles are collected to improve the signal to noise ratio. When a population of dyes

does not diffuse, the sample must be moved after each recovery period in order that

a new spot with a fresh population of dyes may be observed.

The power of the bleaching pulse is chosen to be high enough to destroy the

fluorophores without hurting the cell. The duration of the bleaching pulse is chosen

to be much shorter than the time scale of the diffusion we hope to observe. The probe

power has to be low enough not to cause bleaching during the recovery.

We are interested in the fraction of mobile channels as well as the rate at which

they diffuse. Figure 4.1 illustrates the terminology used to describe the results of a

FRAP experiment. The pre-bleach intensity is the intensity of the fluorescence before

the bleaching pulse. After the bleaching pulse, the fluorescence is decreased. The

bleach depth is the amount of intensity destroyed by the bleaching pulse. The term

“bleach depth” can also refer to the percentage of the pre-bleach intensity removed

by the bleaching pulse. The bleaching pulse is followed by the probe beam during

which the recovery of fluorescence is monitored until the slope of the recovery returns

to zero. The unrecovered fraction is the difference between the pre-bleach intensity

and the intensity at the end of the recovery normalized by the bleach depth. By

40

00 400 800 1200 1600

time (ms)

recovery

bleachingpulse

pre−bleach intensity

time

fluo

resc

ence

inte

nsity

blea

ch d

epth unrecovered fraction

Figure 4.1: Illustration of terminology used in FRAP experiments.

extension, the percent recovery is one minus the unrecovered fraction. The percent

recovery represents the “mobile fraction” of channels.

4.2 Materials and methods

4.2.1 Sample preparation

Initially, oocytes were harvested and injected by our collaborators