, ‘fimw‘

 

 

.J‘ x.
as .6 gfik
13:2... .1
i 2. , .3...

M. .. ‘

vananordnmu‘i
. 4.15.5 of

1A!
my.

a...
any...
:dxmnam1
....:v«.£ .
. 1.1.

mania... .1.
3.1

m. 5 ,3).
4 . an? ‘
hfiw
l‘lliP
v2. 2:9.
‘
a

nmfl! nan...“
it... .25. Zitfilgdawtfi“
3A.”

.3 c u!
.. .1...\72$. . :71, 3....
.1. 51::

1&3...

«flung

:-
1...:

In: .
Lanai.“
(and: 1:)

awn“. run

.533“! ix
.Wr. ulna-H. J. 55:00.:1.
$5.33: I.»

 

 

:.

 

““v\ n.

This is to certify that the
dissertation entitled

Motional and Excitation Transport Dynamics in Restricted
Environments

presented by
Sarah Ann Stevenson

has been accepted towards fulfillment
of the requirements for the

Ph.D. degree in Chemistry

Major/Professor’s Signature

{fig/ac

Date

MSU is an Affirmative Action/Equal Opportunity Institution

 

LIBPARY
ticnigan State
University

 

 

 

 

PLACE IN RETURN BOX to remove this checkout from your record.
TO AVOID FINES return on or before date due.
MAY BE RECALLED with earlier due date if requested.

 

DATE DUE DATE DUE DATE DUE

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

2/05 p:/ClRC/DateDue.indd-p.1

 

MOTIONAL AND EXCITATION TRANSPORT DYNAMICS IN RESTRICTED
ENVIRONMENTS

By

Sarah Ann Stevenson

A DISSERTATION

Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of

DOCTOR OF PHILOSOPHY

Department of Chemistry

2006

ABSTRACT

MOTIONAL AND EXCITATION TRANSPORT DYNAMICS IN RESTRICTED
ENVIRONMENTS

By

Sarah Ann Stevenson

Micelles and vesicles are of interest due to their potential application as
simplistic model biological systems. We are particularly interested in the
interfacial region between the micelle or vesicle polar region and the bulk solution
because this portion of the system mediates exchange processes and
partitioning. To examine the interfacial region of micelle systems, we sequester a
nonpolar optical donor within a micelle and monitor energy transfer from this
molecule to an ionic optical acceptor present in the aqueous phase of the
solution. The time constant for this energy transfer is monitored using time-
correlated single photon counting measurements. Our data indicate that
interactions between the polar head group molecules mediate the efficiency of
energy transfer, and we relate this finding to the presence of the micellar
electrical double layer. The alteration of the ionic strength of the solution will
change the balance of forces that operate within the palisades layer and,
depending on the details of the interactions, the micelle structure may be altered
as a result. We then expand our discussion to the differences between the
interfacial regions of micelle and vesicle systems. These two systems, although

similar, are difficult to compare directly due to the difference in the constituents

which form the structures. We use the molecule decanoic acid and its conjugate
base sodium decanoate, which form micelles or vesicles depending on solution
parameters such as pH, to make this comparison. Our data indicate that,
although these systems share a common constituent, there are measurable
differences between the micellar and vesicular environments. Our findings show
that the environment(s) experienced by chromophores sequestered in the
nonpolar regions of the micelle and vesicle structures are characterized by
significantly different motional constraints because of the different molecular

organization of micelles and vesicles.

To my parents

ACKNOWLEDGMENTS

My first thanks must go to my husband Matt, for his support,
understanding, and willingness to move back to Michigan. My parents, Don and
Nancy Remus, have also been instrumental in teaching me the value of a good
education, as I’m sure was conveyed to them through their own parents. Without
their support throughout my entire education I wouldn’t be where I am today. My
entire family, especially my little brother Jim, has been vital in keeping me in
touch with, or at least offering glimpses of, the “real world”.

There are many other people who contributed to this dissertation in many
ways. I am particularly indebted to my advisor, Gary Blanchard, for his
willingness to help me conquer the steep edge of the learning curve as well as
his encouragement and insight. I would also like to thank my committee
members, Professors Marcos Dantus, Jim McCusker, and David Weliky. Finally, I
would like to offer appreciation for all of my other MSU colleagues and Blanchard
group members past and present for their scientific insight and comic relief. MSU

wouldn’t have been the same without any of you!

TABLE OF CONTENTS

List of Tables ....................................................................................... vii
List of Figures ........................................................................................ x
Chapter 1 - Introduction ......................................................................... 1

Literature Cited ................................................................................. 16

Chapter 2 - Dynamics of 4-Benzylamino-7-nitrobenzofurazan in the 1-
PropanoI/Water Binary Solvent System. Evidence for Composition-Dependent
Solvent Organization

Introduction ..................................................................................... 21

Experimental .................................................................................... 24
Results and Discussion ...................................................................... 25
Conclusions ..................................................................................... 40
Literature Cited ................................................................................. 41

Chapter 3 - Investigating the Micelle-Solution lnterfacial Region Using
Fluorescence Resonance Energy Transfer

Introduction ..................................................................................... 45
Experimental .................................................................................... 48
Results and Discussion ...................................................................... 52
Conclusions ..................................................................................... 77
Literature Cited ................................................................................. 79

Chapter 4 - Investigating Internal Structural Differences Between Micelles and
Unilamellar Vesicles of Decanoic Acid/Sodium Decanoate

Introduction ..................................................................................... 84
Experimental .................................................................................... 88
Results and Discussion ...................................................................... 92
Conclusions ................................................................................... 102
Literature Cited ............................................................................... 103

Chapter 5 - Investigating the Hydrophobic Region of Micelles and Unilamellar
Vesicles Using Fluorescence Resonance Energy Transfer

Introduction ................................................................................... 107
Experimental ................................................................................. 1 1 1
Results and Discussion .................................................................... 115
Conclusions ................................................................................... 132
Literature Cited ............................................................................... 133

vi

Chapter 6 — Conclusions and Future Work

vii

LIST OF TABLES

Table 2.1 - Solvent association calculations for V = mebe + nVsolvent- Values of V

calculated from experimental tog times are shown in the second column, and
from these data we can estimate the number of 1-propanol molecules that
reorient with the 880 chromophore. From this estimate of n and experimental

viscosity data (from Ref 52'5“), we calculate a model TOR ............................... 37

Table 3.1 - Fluorescence lifetime data for perylene sequestered in SDS micelles
as a function of optical acceptor concentration. The data reported here are the
averages of 6 independent determinations, and the uncertainties reported are

:1 o .................................................................................................... 57

Table 3.2 - Fluorescence lifetime data for perylene sequestered in CTAB micelles
as a function of optical acceptor concentration. The data reported here are the
averages of 4 independent determinations, and the uncertainties reported are :1:

1 o ..................................................................................................... 59

Table 3.3 - Equilibrium constant calculations for DSF and R110 in SDS and
CTAB ................................................................................................ 63

Table 3.4 - Rotational diffusion time constants of perylene sequestered in SDS
micelles as a function of optical acceptor concentration. The data reported here
are the averages of 6 independent determinations, and the uncertainties reported
are :t to ............................................................................................. 68

Table 3.5 - Rotational diffusion time constants of perylene sequestered in CTAB
micelles as a function of optical acceptor concentration. The data reported here
are the averages of 4 independent determinations, and the uncertainties reported
are :1: 1o ............................................................................................. 70

Table 5.1 - (a) Fluorescence lifetime data for perylene sequestered in decanoate
vesicles. (b) Fluorescence lifetime data for perylene sequestered in
decanoate/decanoic acid micelles. The data reported here are the averages of

6 independent determinations, and the uncertainties reported are :t to ........... 120

viii

Table 5.2 - (a) Rotational diffusion time constants of perylene sequestered in the
nonpolar region of decanoic acid/decanoate vesicles (b) Rotational diffusion time
constants of perylene sequestered inside decanoate micelles. The data reported
here are the averages of 6 independent determinations, and the uncertainties
reported are 1 1o ................................................................................ 130

LIST OF FIGURES

Figure 1.1 - Micellar Structure and Double Layer .......................................... 4

Figure 1.2 - Schematic of time-correlated single photon counting laser
spectrometer ........................................................................................ 8

Figure 1.3 - Experimental I and l scans for perylene in CTAB, along with the
instrumental response function. These data are typical of those recorded for
reorientation measurements ................................................................... 11

Figure 1.4 - Anisotropy function, R(t), for the data shown in Figure 3. The decay
is fit to the function R(t) = R1(0)exp(t/t) ...................................................... 12

Figure 2.1 - The structure of 4-Benzylamino-7-Nitrobenzylfurazan (BBD) ........ 23

Figure 2.2 - (a) Absorption and emission spectra of BBD as a function of 1-
propanol/water solvent system composition. The absorption and emission
spectra shift to shorter wavelength with increasing 1-propanol concentration.

(b) The Stokes shift of BBD as a function of 1-propanol/water solvent system
composition (5% error) .......................................................................... 26

Figure 2.3 - Permanent dipole moment magnitude and orientation for So (top)
and S1 (bottom) BBD. These calculations were performed at the semi-empirical
level using a PM-3 parameterization. Note that the orientation of the permanent
dipole moment changes little on excitation ................................................. 28

Figure 2.4 - Fluroescence lifetime of BBD as a function of 1-propanol
concentration ....................................................................................... 30

Figure 2.5 - (a) Fluroescence anisotropy decay times as a function of solution 1-
propanol concentration. The calculated model times from Table 1 are plotted for
comparison to the experimental data. Error bars are indicated on individual data
points and are the result of at least six individual determinations of each point.

(D) Solution bulk viscosity of the 1-propanoVwater solvent system as a function of
1-propanol concentration. These data were adapted from ref 52‘“ .................. 34

Figure 3.1 - Structures of the chromophores used in this chapter ................... 49

Figure 3.2 - Cryo-TEM images of SDS (a) and CTAB (b) micelles, showing a
range of sizes less than 100nm diameter .................................................. 51

Figure 3.3 - Normalized absorption and spontaneous emission spectra of
perylene, and absorption spectra of DSF and R110 .................................... 55

Figure 3.4 - Fluorescence lifetimes of perylene sequestered in micelles as a
function of optcal acceptor concentration. (a) SDS micelles, . = DSF, = R110.
(b) CTAB micelles, . = DSF, = R110 ...................................................... 72

Figure 3.5 - Rotational diffusion time constants of perylene sequestered in
micelles as a function of optcal acceptor concentration. (a) SDS micelles, o =
DSF, = R110 (b) CTAB micelles, . = DSF, = R110 ................................ 73

Figure 4.1 - The structures of perylene, sodium decanoate, and decanoic
acid ................................................................................................... 87

Figure 4.2 - (a) TEM image of decanoic acid/decanoate unilamellar vesicles. The
average size is ca. 100 nm diameter. (D) Cryo-TEM image of decanoate micelles
indicating a characteristic diameter of ca. 10 nm ......................................... 90

Figure 4.3 - (a) Experimental l.(t) and Mt) scans for perylene in decanoate
micelles, along with the instrumental response funcation. These data are typical
of those obtained for reorientation measurements. (b) Anisotropy function, R(t),
generated from the data shown in (a). The decay is single exponential and is fit
to the function R(t) = R(O) exp(-t/r) ........................................................... 94

Figure 4.4 - (a) Experimental l.(t) and I l(t) scans for perylene in decanoate
vesicles, along with the instrumental response funcation. These data are typical
of those obtained for reorientation measurements. (b) Anisotropy function, R(t),
generated from the data shown in (a). The decay is single exponential and is fit
to the function R(t) = R(O) exp(-t/r) ........................................................... 96

Figure 4.5 - (a) Normalized absorption and emission spectra of perylene in 100
mM decanoate/decanoic acid micelles (b) Normalized absorption and emission
spectra of perylene in 100 mM decanoate/decanoic acid vesicles .................. 98

Figure 5.1 - The structures of perylene, sodium decanoate, and decanoic
acid ................................................................................................. 110

Figure 5.2 - (a) TEM image of decanoic acid/decanoate unilamellar vesicles. The
average size is ca. 100 nm diameter. (b) Cryo-TEM image of decanoate micelles
indicating a characteristic diameter of ca. 10 nm ....................................... 113

Figure 5.3 - (a) Experimental scans for perylene in decanoate vesicles, along
with the instrumental response function. (b) Experimental scans for perylene in
decanoate vesicles and 10'3 M DSF present. These data are typical of those
obtained for lifetime measurements ........................................................ 118

Figure 5.4 - Experimental scans for perylene in decanoate micelles, along with
the instrumental response function ......................................................... 119

xi

Figure 5.5 - (a) Experimental l.(t) and l 1(t) scans for perylene in decanoate
vesicles, along with the instrumental response funcation. These data are typical
of those obtained for reorientation measurements. (b) Anisotropy function, R(t),
generated from the data shown in (a). The decay is single exponential and is fit
to the function R(t) = R(O) exp(-t/r) ......................................................... 127

Figure 5.6 - (a) Experimental l.(t) and li(t) scans for perylene in decanoate
micelles, along with the instrumental response funcation. These data are typical
of those obtained for reorientation measurements. (b) Anisotropy function, R(t),
generated from the data shown in (a). The decay is single exponential and is fit
to the function R(t) = R(O) exp(-t/r) ......................................................... 128

xii

LIST OF ABBREVIATIONS

TCSPC ............................................ Time Correlated Single Photon Counting
CFD ........................................................... Constant fraction discriminator

TAC ................................................................ Time to amplitude converter
MCP ........................................................................... Microchannel plate
PMT .......................................................................... Photomultiplier tube

MCA ........................................................................ Multichannel analyzer
BBD ...................................................... 4-Benzylamino-7-nitrobenzofurazan
DSF ......................................................................... Disodium Fluoresoein
R110 ................................................................... Rhodamine 110 Chloride
TEM ........................................................ Transmission Electron Microscope
Cryo-TEM .................................. Cryogenic Transmission Electron Microscope

xiii

Chapter 1
INTRODUCTION

Understanding the dynamics of intermolecular interactions in solutions has
been of considerable interest because of the importance of these properties in
mediating a number of different phenomena. Solution phase interactions play an
essential role in chemical separations, catalysis, and several biologically
important processes. In order to gain a better understanding of these processes,
a more complete understanding of the dynamic interactions between the solute
and solvent molecules is necessary. The common underlying issue that has
been most difficult to address is the existence, persistence length and
characteristic time-scale of local heterogeneity.

Amphiphilic molecules are examples of heterogeneous systems that
exhibit self-assembly behavior. Depending on their structural details, these
molecules form bilayer structures, which are the basic component of cell
membranes, and also find use in other applications such as detergents, chemical
separations, and drug delivery. Two subsets of amphiphilic systems that manifest
self-assembly behavior in certain concentration regimes are micelles and
vesicles. Micelles have attracted substantial attention from the chemical
community due mainly to their utility as interfaces between polar and nonpolar
phases. This property has also manifested itself in the use of micelles as
encapsulating and biomimetic systems.“4 Since most of the utility of these

biomimetic structures rests in their ability to exchange species between the

hydrophobic and hydrophilic regions of the systems, it is the interfacial region
which mediates these processes that is of interest.

For ionic micelles, the interfacial region between the micelle and the
aqueous phase is known as the Stern layer. This region contains the ionic
headgroups as well as the majority of the counterions present in the solution,
which causes the micellar surface to resemble an electric double layer.5 As
shown in Figure 1, the Stern layer contains most of the counterions, which are
electrostatically bound to the micelle surface.6 After the Stern layer, a diffuse
layer is present in which electrostatic interactions between solution phase ions
and the micellar surface cause most of the remaining counterions to accumulate
in this region. Following the Stern and diffuse layers is the bulk solution. The role
that the electrostatic double layer plays in mediating interactions between the
polar and nonpolar regions of the micellar systems is not known, and more
insight is needed in order to gain a better understanding of the exchange
processes between micelles and the solution phase.

Like micelles, vesicles are also commonly used as biomimetic systems and for
applications such as drug delivery and immunological studies. Vesicles are
usually prepared from phospholipids, however more simplistic amphiphilic

7-14

compounds have also been shown to form vesicular structures. Monnard ef

al. have recently addressed issues of stability, permeability, and encapsulation of
macromolecules in vesicles prepared from these simplistic fatty acid amphiphilic

molecules.9 These amphiphiles offer advantages over phospholipids, such as

11,13,15-17

their structural simplicity, dynamic behavior, ability to grow upon

13J8

addition of more fatty acid, and autocatalytic or self reproducing

12J9

formation, making them well suited to fundamental examinations aimed at

understanding organization in these dynamic structures. Recent works by
Morigaki et al. and Namani et al. have investigated the sodium
decanoate/decanoic acid fatty acid system and have shown that the same
amphiphilic structures will form micelles or vesicles depending on the pH of the

1020

solution. Since the same constituents form both single and double layer

structures, a direct comparision between micelles and vesicles is possible. We
are particularly interested in the interfacial regions of these structures and any
differences that might exist in their molecular organization.

To elucidate the exchange processes in these model biological systems,
various methods have been used to study micelles, such as small-angle neutron
scattering, NMR, fluorescence spectroscopy, ‘EPR, X-ray scattering, and an
assortment of other techniques.3""2"23 Fluorescence spectroscopy finds wide use
because of its characteristically high sensitivity combined with the range of
chemical properties available with the fluorophores used. For measurements in
heterogeneous systems, it is possible to use fluorescent compounds that will
partition selectively into a given phase, or will reside at the interface between two
phases. This capability, when combined with fluorescence resonance energy

transfer (FRET) measurements, has been used in the study of heterogeneous

2445

systems such as micelles, vesicles, and lipid bilayers. The transfer of energy

O

  

@039

k Diffuse Layer Bulk
Stern Layer

G Water

. Counter-ion

 

Figure 1.1- Micellar Structure and Double Layer

between molecules can occur by a number of means, ranging from dipolar
coupling to collisional quenching. We are concerned in this work with dipolar

coupling because of the characteristic length scale over which it operates. The

transfer of energy by dipolar coupling has been described by Forster“,

_ _ 6
k —"2kD 8.8x1017I8A0/If0f(v)d17=fl£[%] 1.1

 

DA: n4n6 n4

where kDA is the rate constant for the transfer of energy between donor and

acceptor. The x term is a function of the relative orientation of the donor and

acceptor transition dipole moments, and for our experimental conditions,

«2 >= 213. The term krD is the donor radiative decay rate constant in the

absence of optical acceptor, n is the refractive index and R is the intermolecular
D-A distance. The integral term is a measure of the spectral overlap between the
emission spectrum of the optical donor and the absorption spectrum of the
optical acceptor. The spectral overlap term, combined with the several constants

on the right hand side of Equation 1.1 is typically referred to as the critical or

Forster radius, R0, a quantity which serves as an effective gauge of the operating
distance of this dipolar coupling process. While the term R0 will vary for each

donor-acceptor pair, for most organic chromophore pairs, R0 ~ 50 Aim”

The topology of these systems can significantly influence the efficiency of
energy transfer since the dipolar coupling process that mediates energy transfer
depends on both the average distance and orientation of the molecules involved.

Energy transfer measurements can be used as a “spectroscopic ruler" to

estimate average distances between optical donor and acceptor, and thus we
can gain information on the extent to which micelle and acceptor interact.
Micellar and vesicular systems are ideal candidates for energy transfer
experiments, and previous work with micelles has examined energy transfer
between two molecules sequestered inside a micelle, between a molecule inside

the micelle and a molecule that is part of the surfactant, and between two

26.232.35.49

molecules on the surface of the micelle. Most of these studies have

focused on energy transfer between molecules in the same region of the micelle.
We are particularly interested in the interfacial region of micelles, and
understanding the intermolecular dynamics in micellar systems will hopefully
shed some light on more complex biomimetic systems. Previous work with
vesicle systems has explored different aspects of vesicle structure and behavior.
Earlier work by Fung and Stryer monitored membrane fusion between vesicles
labeled with fluorescent probes and unlabeled vesicles."o For simpler single

chain amphiphilic vesicles, Chen utilized FRET, or Fluorescence Resonance
Energy Transfer, to examine the kinetics of fatty acid vesicle growth.13 Several

researchers, such as Dobretsov et al. and van Zandvoort ef al. have used FRET

measurements to determine the location of various fluorescent probes in lipid

51,52

membranes. While all of these studies have lead to greater understanding of

vesicle systems, a direct comparison between micellar and vesicular structures
remains to be made.
To probe the local environments of the micelle and vesicle systems, we

use fluorescence lifetime and depolarization measurements. The picosecond

time-correlated single photon counting laser spectrometer used for all of the

fluorescence lifetime and depolarization experiments has been described
previously53 and we recap a brief outline of its performance features here. A

graphical depiction of the instrument is offered in Figure 1.2. The source laser is
a mode locked CW Nd:YAG laser (Coherent Antares 76s), that produces 100 ps
1064 nm pulses at a 76 MHz repetition rate. The 1064-nm pulse train is
frequency tripled to produce ~1.3 W average power at 354.7 nm for all
experiments. The third harmonic (354.7 nm light) of the Nd:YAG laser is used to
excite a Coherent 702-2 cavity dumped dye laser. The dye laser is cavity
dumped at a repetition rate of 4 MHz and can generate pulses ranging from ~
550 -1000 nm depending on the optics used. For these experiments, we
produced output using Stilbene 420 laser dye (Exciton) to produce 420 nm light
for experiments involving perylene and 460 nm light for 4-bezylamino-7-
nitrobenzofurazan (BBD) experiments. Excitation power at the sample was s 1
mW average power for all measurements. The output from the dye laser is
divided and half of the light is sent through the reference channel while the other
half travels to the sample holder. The reference channel was delayed optically

and detected using an in-house built fiber optic delay line. Fluorescence was

collected at at 0°, 54.70 and 90° with respect to the vertically polarized excitation

pulse with the polarization being selected by a Glan-Taylor prism. The
polarization of this selected light is then scrambled and a subtractive double
monochromator (American Holographic DB10-S) is used for wavelength

selection. A subtractive double monochromator is utilized to reduce the effects of

 
 
 
 
      
  

3w

cavity dumped dye laser

 

  
       
 
 

A

l

r
.I

 

(1mm

neutral density
filter

.x" .-' ‘

GfT monochromator
pnsm

Figure 1.2 - Schematic of time-correlated single photon counting

laser spectrometer.

fiber optic

'* g switchable

delay

MCP PMT

 

 

dispersion-induced time broadening of the collected light.54 Fluorescence signals
were detected using a Hamamatsu R3809U microchannel plate photomultiplier
tube that has a rise time of 156 ps and a transit time spread of 42 ps FWHM. The
signal was then sent to a Tennelec 454 quad constant fraction discriminator
where a Tennelec 864 time-to-amplitude converter and biased amplifier is used
for signal processing. For most TCSPC measurements, the systems are typically
operated in reverse mode to avoid temporal distortions caused by electronic
dead time limitations.”55 The remaining half of the dye laser output is coupled
into an optical fiber and then routed to a custom made fiber optic delay line”,
which allows a range of lifetimes from microseconds to picoseconds. A
Hamamatsu S2381 avalanche photodiode collects the optically delayed laser
output and is input to the CPU. The signal from the reference channel is delayed
electronically (T ennele 412A) and travels to the stop channel of the TAC. TAC
output counts are observed using a Tektronix 100-MHz oscilloscope and are sent

to a Tennelec PCA-11 multichannel analyzer for collection. The fluorescence

collected at 54.7° polarization provides fluorescence lifetime data by eliminating

molecular contributions to the signal while 0° and 90° polarizations are of

interest for rotational diffusion measurements. Data were acquired and the

collection wavelength and polarization selector were computer controlled using a

program written with LabVlEW® v. 7.1 and a Newport PMC200-P programmable

motion controller. Lifetimes range from ~800 ps to ~6000 ps, while an instrument

response function of typically 35 ps fwhm is observed for this system. We did not
deconvolute the instrument response function from the fluorescence transients.

To observe the interactions of solvent molecules in the solution phase we
utilize two complementary techniques; fluorescence lifetime and depolarization
measurements. Fluorescence lifetime data provide information about the local
environment of the probe molecule as well as the extent of fluorescence
resonance energy transfer while fluorescence depolarization measurements offer
insight into the microenvironments that exist within the heterogeneous systems.
Lifetime measurements give us a qualitative picture of the interfacial region while
reorientation data provide us with quantitative information regarding the local
viscosity of the nonpolar regions of micelle and vesicle systems.

In rotational diffusion measurements, the probe molecule rotates in
solution and sweeps out a particular volume. The volume that the probe sweeps
out depends on the size and shape of the probe molecule as well as the
properties of the surrounding solvent. By examining the rotational dynamics of a
probe molecule in solution, we can gain insight into the local environment of the
probe molecule. Reorientation dynamics have been used extensively to
investigate local organization in solutions because of the well developed

theoretical framework for interpretation of the data and the relatively high

56-60

information content of the experimental signals. Using vertically polarized

light we can preferentially excite those probe molecules whose transition dipole

moments are parallel to the excitation. This induced anisotropy decays over time,

10

2000

1000

 

 

Signal Intensity (a.u.)

 

 

 

 

6000 12000 18000
Delay Time (ps)

Figure 1.3 — Experimental l and LI. scans for perylene In CT AB, along with
the Instrumental response unction. These data are typical of those
recorded for reorientation measurements.

11

 

 

 

 

 

 

 

 

 

 

 

0.10 I)
l)"
1% I ‘
g 0.051 ‘b I
§ “NW
é Il ,
0.011 "MI ”WI, ‘I'UIIIIIIIIQV l
0 200 400 5'00 7 W—

Delay Time (ps)

Figure 1.4 - Anisotropy function, R(t), for the data shown In Figure 1.3. The
decay is fit to the function R(t) = R(0)exp(-tlr)

12

providing information on the rotational diffusion dynamics. The induced
orientational anisotropy function, R(t), is shown in equation 1.2 and is obtained
by combining the data collected parallel and perpendicular to the excitation

polarization,

[um-Mt)

R(t): I"(t)+2li(f)

1.2

 

where I"(t) and ll(t) are the polarized emission intensities. A representative
sample data set is presented in Figure 1.3. The chemical information contained
in these data is in the decay time constant(s) of R(t), and its zero-time value,

R(O). The theory has been well established by Chuang and Eisenthal, who
calculate that R(t) can decay with up to five exponential terms.56 For most

chromophores, one or two exponentials are typically observed and the
functionality of the decay depends on the orientation of the chromophore
transition dipole moments relative to the rotational motion axes and the shape of
the ellipsoid swept out by the rotating moiety. Several physical models have been
presented to interpret the experimental data. The choice of model will determine
the physical and chemical significance of the decay time constant(s) extracted
from the R(t) data. For chromophores tethered to a substantially larger entity,
such as a protein, or imbedded in a mono- or bilayer structure, the hindered rotor
model is suitable. In this model the chromophore is attached to the larger
assembly at one point within the system, and its motion is considered in the
context of the conic volume it sweeps out. For chromophores that are able to

rotate freely in solution, the modified Debye-Stokes-Einstein (DSE) equation is

13

the preferred model. The modified Debye—Stokes-Einstein (DSE) equation is

given by Eq. 1.2,61

_ 77Vf

Tor — kBTS 1 .3

where n is the solvent viscosity, V is the solute hydrodynamic volume, T is the

temperature, k3 is the Boltzmann constant, f is the frictional interaction term,°°

and Sis the solute molecular shape factor.62 The DSE model is limited in the

sense that it does not account for the molecular nature of the solvent, yet for
many systems it provides remarkably close agreement with experimental data.
We make use of this model throughout this dissertation to help in understanding
the behavior of chromophores sequestered within the micelle and vesicle
structures we consider here.

These fluorescence measurements can be used to gain insight about the
model biological systems that we have choosen to investigate. To utilize these
measurements, we use fluorescent probe molecules to observe changes in local
environments. Perylene is an extensively studied hydrophobic chromophore that
is known to position itself in nonpolar environments. For the heterogeneous
micellar and vesicular systems that we would like to observe, we are able to
sequester perylene in the nonpolar regions of these systems.

Chapter 2 of this dissertation examines the micelle-solution interface using
fluorescence resonance energy transfer. We have found that the presence of

counterions in ionic surfactants mediate the excitation transport between a probe

14

sequestered inside a micelle and one present in the aqueous phase of the
solution.

Chapter 3 deals with the dynamics of 4-benzylamino-7-nitrobenzofurazan
in the 1-propanol-water binary solvent system. It is clear that in the 1-

propanoVwater solvent system there is discernible microscopic solvent
heterogeneity, with the details of this transient solution phase structure
depending sensitively on the composition of the solvent system.

Chapters 4 and 5 examine the differences between micelles and vesicles.
We look at the different local environments of decanoate/decanoic acid micelles
and vesicles in Chapter 4. In Chapter 5, we look at the communication between a
probe molecule sequestered in the nonpolar region of the micelle or vesicle
structure and a probe in the solution phase.

Chapter 6 provides some final conclusions as well as possible
suggestions for future experiments to further examine the sodium

decanoate/decanoic acid system.

15

Literature Cited

(1) lnada, A.; Nakamura, Y.; Morita, Y. Chemical & Pharmaceutical
Bulletin 1988, 36, 462.

(2) lkura, T.; Go, N.; lnagaki, F. Proteins: Structure, Function, and
Genetics 1991, 9, 81.

(3) Hwang, P. M.; Vogel, H. J. Biochemistry and Cell Biology 1998, 76,
235.

(4) Henry, G. D.; Sykes, B. D. Methods in enzymology FIELD
Publication Date: 1994, 239, 515.

(5) RangeI-Yagui, C. 0.; Pessoa, A., Jr.; Tavares, L. C. Journal of
Pharmacy & Pharmaceutical Sciences 2005, 8, 147.

(6) lsraelachvili, J. lnterrnolecular & Surface Forces, second edition
ed.; Academic Press: San Diego, 1992.

(7) Gebicki, J. M.; Hicks, M. Nature (London, United Kingdom) 1973,
243, 232.

(8) Hargreaves, W. R.; Deamer, D. W. BiochemistryI978, 17, 3759.

(9) Monnard, P.-A.; Deamer, D. W. Methods in Enzymology 2003, 372,
133.

(10) Morigaki, K.; Walde, P.; Misran, M.; Robinson, B. H. Colloids and
Surfaces, A: Physicochemical and Engineering Aspects 2003, 213, 37.

(11) Sacerdote, M. G.; Szostak, J. W. Proceedings of the National
Academy of Sciences of the United States of America 2005, 102, 6004.

(12) Walde, P.; Wick, R.; Fresta, M.; Mangone, A.; Luisi, P. L. Journal of
the American Chemical Society 1994, 116, 11649.

16

(13) Chen, I. A.; Szostak, J. W. Biophysical Joumal2004, 87, 988.

(14) Cistola, D. P.; Hamilton, J. A.; Jackson, 0.; Small, D. M.
Biochemistry 1988, 27, 1881 .

(15) Kamp, F.; Zakim, D.; Zhang, F.; Noy, N.; Hamilton, J. A.
Biochemistry 1995, 34, 1 1928.

(16) Zhang, F.; Kamp, F.; Hamilton, J. A. Biochemistry 1996, 35, 16055.

(17) Kleinfeld, A. M.; Chu, P.; Romero, C. Biochemistry 1997, 36,
14146.

(18) Hanczyc, M. M.; Fujikawa, S. M.; Szostak, J. W. Science
(Washington, DC, United States) 2003, 302, 618.

(19) Oberholzer, T.; Wick, R.; Luisi, P. L.; Biebricher, C. K. Biochemical
and Biophysical Research Communications 1995, 207, 250.

(20) Namani, T.; Walde, P. Langmuir 2005, 21, 6210.

(21) Griffiths, P. C.; Cheung, A. Y. F.; Finney, G. J.; Farley, C.; Pitt, A.
R.; Howe, A. M.; King, S. M.; Heenan, R. K.; Bales, B. L. Langmuir2002,
18, 1065.

(22) ltri, R.; Amaral, L. Q. Journal of Physical Chemistry 1991, 95, 423.
(23) Paradies, H. H. Colloids and Surfaces 1983, 6, 405.

(24) Barzykin, A. V.; Tachiya, M. Journal of Chemical Physics 1995,
102, 3146.

(25) Birch, D. J. S.; Suhling, K.; Holmes, A. S.; Salthammer, T.; lmhof,
R. E. Pure and Applied Chemistry 1993, 65, 1687.

(26) Choi, K. J.; Turkevich, L. A.; Loza, R. Journal of Physical Chemistry
1988. 92, 2248.

17

(27) de Oliveira, H. P. M.; Gehlen, M. H. Chemical Physics 2003, 290,
85.

(28) DelaCruz, J. L.; Blanchard, G. J. Journal of Physical Chemistry B
2003, 107, 7102.

(29) Dutt, G. B. Journal of Physical Chemistry B 2002, 106, 7398.

(30) Ediger, M. D.; Domingue, R. P.; Fayer, M. D. Journal of Chemical
Physics 1984, 80, 1246.

(31) Ediger, M. D.; Fayer, M. D. Journal of Chemical Physics 1983, 78,
2518.

(32) Johansson, L. B. A.; Niemi, A. Journal of Physical Chemistry 1987,
91. 3020.

(33) Kelepouris, l..; Blanchard, G. J. Journal of Physical Chemistry B
2002, 106.6600.

(34) Kelepouris, L.; Blanchard, G. J. Journal of Physical Chemistry B
2003, 107, 1079.

(35) Kenney-Wallace, G. A.; Flint, J. H.; Wallace, S. C. Chemical
Physics Letters 1975, 32, 71.

(36) Klein, U. K. A.; Haar, H. P. Chemical Physics Letters 1978, 58, 531.

(37) Maiti, N. C.; Krishna, M. M. G.; Britto, P. J.; Periasamy, N. Journal
of Physical Chemistry B 1997, 101, 1 1051.

(38) Maiti, N. C.; Mazumdar, S.; Periasamy, N. Journal of Physical
Chemistry 1995, 99, 10708.

(39) Mattjus, P.; Molotkovsky, J. G.; Smaby, J. M.; Brown, R. E.
Analytical Biochemistry 1999, 268, 297.

18

(40) Menger, F. M. Accounts of Chemical Research 1979, 12, 111.

(41) Quitevis, E. L.; Marcus, A. H.; Fayer, M. D. Journal of Physical
Chemistry 1993, 97, 5762.

(42) Rao, U. R. K.; Manohar, C.; Valaulikar, B. S.; lyer, R. M. Journal of
Physical Chemistry 1987, 91, 3286.

(43) Saha, D. 0.; Ray, K.; Misra, T. N. Spectrochimica Acta, Part A:
Molecular and Biomolecular Spectroscopy 2000, 56A, 797.

(44) Stryer, L. Annual Review of Biochemistry 1978, 47, 819.

(45) Wistrom, C. A.; Jones, G. M.; Tobias, P. S.; Sklar, L. A. Biophysical
Journal 1996, 70, 988.

(46) Forster, T. Ann. Physik. 1948, 2, 55.

(47) lshii, Y.; Yoshida, T.; Funatsu, T.; Wazawa, T.; Yanagida, T.
Chemical Physics 1999, 24 7, 163.

(48) Hiller David, A.; Fogg Jonathan, M.; Martin Amy, M.; Beechem
Joseph, M.; Reich Norbert, 0.; Perona John, J. Biochemistry FIELD
Publication Date:2003, 42, 14375.

(49) Marcus, A. H.; Diachun, N. A.; Fayer, M. D. Journal of Physical
Chemistry 1992, 96, 8930.

(50) Fung, B. K.-K.; Stryer, L. Biochemistry1978, 17,5241.

(51) Dobretsov, G. E.; Kurek, N. K.; Makhov, V. N.; Syreishchikova, T.
I.; Yakimenko, M. N. Journal of Biochemical and Biophysical Methods
1989, 19, 259.

(52) van Zandvoort, M. A. M. J.; Gerritsen, H. C.; van Ginkel, G.; Levine,
Y. K.; Tarroni, R.; Zannoni, C. Journal of Physical Chemistry B 1997, 101,
4149.

19

(53) DeWitt, L.; Blanchard, G. J.; LeGoff, E.; Benz, M. E.; Liao, J. H.;
Kanatzidis, M. G. Journal of the American Chemical Society 1993, 115,
12158.

(54) Holtom, G. R. Proc. SPIE 1990, 1204, 2.

(55) Haugen, G. R.; Wallin, B. W.; Lytle, F. E. Rev. Sci. Instrum. 1979,
50, 64.

(56) Chuang, T. J.; Eisenthal, K. B. Journal of Chemical Physics 1972,
57, 5094.

(57) Klein, U. K. A.; Haar, H. P. Chemical Physics Letters 1979, 63, 40.
(58) Labhart, H.; Pantke, E. R. Chemical Physics Letters 1973, 23, 482.

(59) Hu, C.-M.; Zwanzig, R. Journal of Chemical Physics 1974, 60,
4354.

(60) Youngren, G. K.; Acrivos, A. Journal of Chemical Physics 1975, 63,
3846.

(61) Debye, P. Polar Molecules. ln Chemical Catalog Co. New York,
1929.

(62) Perrin, F. Journal de Physique et Ie Radium 1936, 7, 1.

20

Chapter 2

DYNAMICS OF 4-BENZYLAMINO-7-NITROBENZOFURAZAN IN THE 1-
PROPANOL-WATER BINARY SOLVENT SYSTEM. EVIDENCE FOR
COMPOSITION-DEPENDENT SOLVENT ORGANIZATION
Introduction
The interactions between dissimilar molecules in the solution phase have
been the subject of a great deal of study because of the importance of such
interactions in processes that include chemical separations, synthesis and
heterogeneous and homogeneous catalysis. In all of this work, the common
underlying issue that has been most difficult to address is the existence,
persistence length and characteristic time-scale of local heterogeneity. We have
been investigating micellar systems "5 and have found that the composition of
the solutions from which the micelles are formed can have an influence on the
resulting micelles. In an effort to understand these effects and in a larger sense
to determine whether or not solution phase heterogeneity plays a role in the
solutions we use, we have studied the lifetime and motional dynamics of a
chromophore dissolved in a series of 1-propanoVwater binary solvent systems.
A variety of spectroscopic methods have been utilized to probe solution

phase systems, including vibrational relaxation?” fluorescence lifetime, and
molecular reorientation measurements.5"7'3‘. In this study, we have chosen to
use time-resolved fluorescence lifetime and anisotropy measurements of 4-
Benzylamino—7-Nitrobenzofurazan (BBD) because of the properties of this
chromophore and the wide use of its structural derivatives in the study of micellar

and lipid bilayer structures. We use rotational diffusion due to the relatively high

21

information content of the signal and the established theoretical framework that is
available for interpretation of the experimental data?"39 While the interpretation
of fluorescence lifetime data is less straightforward and more system-dependent,
for the chromophore we use here, there is an established dependence of its
fluorescence lifetime on the “polarity" of the local environment. The chromophore
that we have chosen for this work (BBD, Figure 10) is a member of a family of
chromophores with the 7-nitrobenz-2-oxa-1,3-diazol-4-yl (NBD) group. These
chromophores are known to exhibit absorption and emission band positions that
are sensitive to solvent polarity, causing them to be used widely as fluorescent
probes in biological and model membrane systems.”43 Shorter fluorescence
lifetimes are observed with more polar solvents, making these probes well suited
for studies of binary solvent systems. We have investigated the 1-propanol/water
solvent system to understand whether or not there is discernible local
heterogeneity in this system. Our data show that there is indeed local
heterogeneity in this binary solvent system, and that the characteristic
persistence time of this transient organization lies in the time window between
the chromophore reorientation time (ca. 100 ps) and its fluorescence lifetime
(several ns). Our reorientation data do not follow the simple viscosity-
dependence predicted by the Debye-Stokes-Einstein (DSE) model (vide infra)
and the composition of the solvent system modulates the strength of solvent-
solute interactions measurably. We find that for the 1-propanoVwater binary
system, there is a range of propanol concentrations for which the solvent-solvent

interactions are strong enough to alter the chromophore interactions with the

22

solvent system. The range of 1-propanol concentrations for which this occurs
coincides with the concentration range for which 1-propanol and water are known
to form an azeotrope. These findings shed new light on the heterogeneity of the

local environment in binary solvent systems.

Figure 2.1 - The structure of 4-Benzylamlno-7-Nitrobenzylfurazan (BBD).

23

 

Experimental

Materials. 4-Bezylamino-7-nitrobenzofurazan (BBD) was obtained from
Sigma-Aldrich, inc. and used as received. 1-Propanol was acquired from CCl,
Inc., and used as received. Water was distilled in-house. The BBD
concentration for all solutions was 10 uM. Solution temperatures were
maintained at 298 10.5 K (Neslab EX100—DD) for all time-resolved fluorescence
lifetime and anisotropy measurements.

Time-Correlated Single Photon Counting Measurements. Fluorescence
lifetime and anisotropy measurements of BBD in a series of 1-propanol/water
solutions were made using a time-correlated single photon counting (T CSPC)
instrument that has been described in detail previously,“ and a more detailed
description can be seen in Chapter One. The excitation for the BBD experiments
is 460 nm, and fluorescence is collected at 490 nm.

Steady-State Spectroscopy. Absorption spectra were recorded with 1 nm
spectral resolution using a Cary model 300 double-beam UV-visible absorption
spectrometer. Spontaneous emission spectra were acquired with a Spex
Fluorolog 3 spectrometer, with both excitation and emission monochromators set
to 3 nm resolution.

Semi-empirical calculations. The permanent dipole moment of BBD was
calculated for the ground state (So) and first excited electronic state (S) using

Hyperchem" v. 6.0, with PM-3 parameterization.45

24

Results and Discussion

We have been involved in the study of micellar systems using optical
spectroscopic methods. Such studies require the incorporation of fluorescent
molecules into micelles, a task which is not always straightforward. Owing to the
typically limited solubility of fluorescent probe molecules in aqueous solutions,
concentrated chromophore solutions made with an alcohol cosolvent are
sometimes used. Even with the presence of a small amount of an alcohol in the
micellar solution, there is a question of how the micelles are altered by its
presence, and this is an issue that we have addressed experimentally.‘ In this
work we are concerned with the intrinsic solution phase heterogeneity that
attends binary 1-propanoVwater solvent systems. If there is, in fact, detectable
solution phase heterogeneity in alcohol/water binary systems, such transient
solution phase organization could give rise to unexpected partitioning
phenomena in multi-component solutions in which micelles are formed. Our
fluorescence lifetime and anisotropy measurements point to local heterogeneity
in 1-propanoVwater solutions, with a characteristic persistence time on the sub-
nanosecond timescale. We find that the 1-propanoI/water system interacts with
the chromophore BBD in a manner that depends sensitively on the composition
of the solvent system, with the nominally weakest BBD-solvent interactions
occurring for solvent ratios close to those for which 1-propanol and water form
azeotropes.

Steady State Spectroscopy. As a first step in the characterization of BBD

in 1-propanoVwater solvent systems, we consider the solvent-dependence of the

25

 

0.8 -

0.6 -

0.4 ~

Linear Response (normalized)

 

0.0 -

 

1 5000 20000 25000
Wavenumber (cm'1)

3200

3100

V I f I j

 

 

 

 

 

 

 

Stokes shift (cm'1)
fl
I
r
1

 

2600 L J“ L " 4- y = 2.82 x + 2603
500

 

 

2300 ° 1 I n I 1 l l I 1 l
0 20 40 60 . 80 100

solution composition (wt-% 1-propanol)

Figure 2.2 - (a) Absorption and emission spectra of BBD as a function of 1-
propanol/water solvent system composition. The absorption and emission
spectra shift to shorter wavelength with increasing 1-propanol
concentration. (b) The Stokes shift of BBD as a function of 1-
propanol/water solvent system composition (5% error).

26

chromophore steady state absorption and emission spectra (Figure 2.2). The
absorption maximum of BBD is shifted to shorter wavelengths with increasing 1-
propanol concentration. This is an expected result since it is known that as the
polarity of the solution increases the absorption maximum shifts to the red. While
the physical origin of the spectral band position dependence on solvent polarity
remains to be elucidated for this chromophore, these steady state data
demonstrate that, on average, as the 1-propanol content of the solvent system is
increased, the chromophore senses this change in a proportional manner. This
is not a surprising result, owing to the expected transient nature of local
organization in the solution phase, and these data are consistent with the
fluorescence lifetime data (vide infra). Since the ground and excited state dipole
moments are not being perturbed, the steady state data can be thought of in the
context of simple energy level shifts.

Fluorescence lifetime measurements. Fluorescence lifetime data are
useful for a variety of reasons. For some chromophores, such as BBD, the
fluorescence lifetime is related to the polarity of the medium, analogous to the
relationship seen for the BBD steady state spectra and solvent polarity. To the
best of our knowledge, there is not a fundamental explanation for the relationship
between solvent properties and BBD spectroscopic response, but there are still
several interesting points that can be discerned from these data.

For many solvatochromic chromophores, the Stokes shift and emission
band positions are related to the ability of the solvent medium to stabilize the

chromophore excited electronic state. We observe for BBD that both the

27

N/ \N
\ /
h 4 N02
So
11:89 D
o
N/ \N
\ /
b A O
.1' N 2
81
u=10.9

Figure 2.3 - Permanent dipole moment magnitude and orientation for So
(top) and 81 (bottom) BBD. These calculations were performed at the semi-
empirical level using a PM-3 parameterization. Note that the orientation of

the permanent dipole moment changes little on excitation.

28

 

absorption and emission bands blue-shift as the polarity of the solvent system is
reduced, while the Stokes shift remains constant (Figs. 2.2). These data suggest
that both the So and S1 electronic states of this chromophore are relatively polar,
with the S1 state being more polar than the So. Specifically, there appears not to
be any site-specific solvent-solute interaction that gives rise to the preferential
stabilization of either the S1 or So electronic states. This interpretation is
supported by semi-empirical calculations of BBD (Figure 2.3) indicating the So
dipole moment to be 8.9 D and the S1 dipole moment to be 10.9 D. Interestingly,
the S1 lifetime increases as the polarity of the solvent decreases, but in all cases
decays as a single exponential (Figure 2.4). The S1 lifetime for any chromophore
can be modulated by a number of factors, including the ability of the solvent
medium to allow a relatively rigid (planar) chromophore. Clearly the data we
present here do not provide sufficient information to elucidate the basis for the
dependence of the fluorescence lifetime on the solvent system polarity, but we
do note that, even if there is discernible heterogeneity in the solvent system, this
structural heterogeneity is effectively averaged over the lifetime of the S1 state of
BBD, thereby placing an upper bound on the persistence time of any solvent
system transient organization (vide infra). With this information in hand, we turn
to a discussion of the fluorescence anisotropy data.

Molecular Reorientation. For the systems under study here, fluorescence
anisotropy data are accounted for in the context of molecular rotational motion.
We obtain fluorescence depolarization data experimentally by exciting BBD with

a vertically polarized light pulse and collecting the emission decay transients

29

 

 

7000
O
6000
8
E) 5000 O
'8 .
5&1 4000
3 e
5 o
8 3000 e
2 e
8 e
'7" 2000
O
O
1000
20 4O 60 80 100

 

solution composition (wt-% l-propanol)

Figure 2.4 - Fluroescence lifetime of BBD as a function of 1-propanol
concentration.

30

polarized parallel and perpendicular to the excitation polarization. These data

are combined to produce the induced orientational anisotropy function, R(t)

rum—1,0)

R r =
U 1,,(z)+21,(t)

(1)

where l"(t) and lj(t) are the polarized emission intensities. The chemical
information contained in these data resides in the decay time constant(s) of R(t),
and its zero-time value, R(O). There is a well established body of theory for the
interpretation of R(t)”? In principle, R(t) can decay with up to five exponential
terms, but for most chromophores Chuang and Eisenthal’s theorys5 reduces to
either one or two exponential decay components, depending on the shape of the
ellipsoid swept out by the rotating moiety and the orientation of the chromophore
transition dipole moments relative to the rotational motion axes. Experimentally,
the most common result is the recovery of a single exponential decay component
in R(t), and that is the functional form of the data we have acquired for BBD in 1-
propanoVwater. The functional form of R(t) allows the interpretation of our data
within the framework of the modified Debye-Stokes-Einstein model.

The modified Debye-Stokes-Einstein (DSE) equation is given by Eq. 233'37

= 77er
Tor kBTS (2)

where n is the solvent viscosity, V is the solute hydrodynamic volume“, T is the

temperature, k3 is the Boltzmann constant, f is the frictional interaction term“,

and Sis the solute molecular shape factor”. The DSE model is limited in the

sense that it does not account for the molecular nature of the solvent, yet for

31

 

many systems it provides remarkably close agreement with experimental data.
The success of this model lies in the fact that the recovered time constant
averages over the many fast molecular interactions that are collectively
responsible for solvation. It is only when the characteristic time constant for the
molecular solvent-solute interactions becomes on the order of the reorientation
time constant that one observes a substantial contribution from discrete
intermolecular interactions. When experimental values diverge from the modified
DSE model predictions, the discrepancy between experiment and model is
typically attributed to deviations in the solvent viscosity, solute hydrodynamic
volume, frictional boundary condition, or solute shape factor. Given the
experimental information available in most cases, it is not possible to clearly
distinguish which of these quantities accounts for experimental behavior.
Solvent-solute interactions dominated by frictional interactions or a discrepancy
between the microscopic and bulk solvent viscosity usually are taken to account
for deviations in the trends predicted by DSE, whereas non-monotonic variations
in Ton with solvent system properties such as composition or bulk viscosity
typically imply an important role for molecular-scale processes. The shape of the
probe molecule is, in general, predicted to be ellipsoidal, with the shape of the
volume swept out during reorientation being described by Perrin’s equations”. In
many cases, especially where single exponential anisotropy decays are
recovered experimentally, the actual volume swept out by the reorienting moiety
is not simply that of the probe molecule. The hydrodynamic volume of the

reorienting moiety can deviate from model predictions due to preferential,

32

relatively long-lived solvent-solute interactions (e.g. H-bonding), and there have
been several accounts of such behavior in binary solvent systems.“-"51

We recover single exponential anisotropy decays for BBD in all 1-
propanoVwater solvent systems. Our data are consistent with BBD reorienting as
a prolate rotor, assuming the S1 ._ So transition moment lies in the chromophore
rt-system plane along the long axis. We recognize that the orientation of the
transition dipole moment may deviate slightly from this geometric axis clue to the
asymmetric substitution of the chromophore 6-membered ring, but such an effect
does not bear on the substance of our findings. We calculate the hydrodynamic
volume of BBD to be 215 A3,“ T=300 K, and i=1 (stick limit). We have not
reported a value for S because of the uncertainty in the volume swept out by the
reorienting chromophore because of the bulky amino-methylphenyl substituent.
Based on these quantities and using Eq. 2, we calculate Ton/n = 51.9 ps/cP in the
stick limit. We show in Figure 14a the dependence of the experimental
reorientation time on the solvent system composition. These data do not follow a
trend that is monotonic with respect to the solvent system bulk composition, and
it is this point which requires a more detailed treatment.

As noted above, trends in reorientation data that scale linearly with
solution viscosity can indicate molecular motion being mediated by processes
that are averaged over a large number of molecular-scale events. To address
whether or not the reorientation data we report here do, in fact, vary
monotonically with solvent viscosity, we must understand how the viscosity of the

1-propanol/water system depends on composition. These data have been

33

N
to
DD

 

 

732 . 3 3
8 18 o O
u 16 ’ 9 . '
'E l . . . O g
c:
.o 12 e
3 1
8 8 ‘
'E‘.
o
8 4
20 4O 6O 80 100
solution composition (wt-% l-propanol)

22
122 b
3‘ 18
“a; 1
' 1
LL]
V1 12
g 1
c0 8
'8‘
'3 4
° 20 4O 6O 80 100

solution composition (wt-% l-propanol)

Figure 2.5 - (a) Fluroescence anisotropy decay times as a function of
solution 1-propanol concentration. The calculated model times from Table
1 are plotted for comparison to the experimental data. Error bars are
indicated on Individual data points and are the result of at least six
Individual determinations of each point. (b) Solution bulk viscosity of the
1-propanollwater solvent system as a function of 1-propanol concentration.
These data were adapted from ref 52'“.

tabulated in the literature,“""'54 and there is a nonlinear dependence of solution

viscosity on solution composition. Using the DSE model parameters given above
and the literature data on 1-propanoVwater viscosity, we show the calculated Ton
times in Figure 2.5b. Clearly the calculated results do not agree well with the
experimental data. At high 1-propanol concentrations, the experimental TOR

values are much larger than the calculated values, and for 1-propanol
concentrations in the region of 30%, there is an anomalous spike in the
experimental reorientation data which is not reflected in the calculations. The
experimental reorientation data cannot be understood wholly in the context of
solvent properties, and the physicochemical phenomena that give rise to these
data operate on a time scale commensurate with the measured anisotropy decay
time(s).

The hydrodynamic volume of BBD is 215 A3 using Edward’s model. This
model is well established and yields results that are consistent with the
reorientation dynamics of many chromophores in neat solvents. The
experimental, non-monotonic deviations from the DSE model predictions are not
associated the calculated volume of BBD, but the total volume of the reorienting
moiety can vary with solution composition. It is possible that one or more solvent
molecules (presumably propanol) are associated with the probe molecule,
yielding a reorienting species that has a larger hydrodynamic volume than that of
the chromophore alone. In this case we describe V = V339 + nvmm, where V is
the total hydrodynamic volume of the reorienting entity, V339 = 215 A3, V5,...“ is

the hydrodynamic volume of a solvent (1 -propanol) molecule, and n is the

35

number of solvent molecules that are closely associated with BBD. Using
Edward’s method, we calculate VW = 70 A3 for 1-propanol, and within the
context of this explanation, the added solvent volume required for consistency of
the model with the data is n = 1 for low 1-propanol concentrations, reducing to no
added solvent volume for solution compositions where 1-propanol and water are
known to form an azeotrope. then increasing to a high value of n = 3 at high 1-
propanol concentrations (see Table 2.1). We note that the agreement between
this model and the experimental data is not exact, but rather produces good
qualitative agreement. We believe that the discrepancies between model and
data lie in the fact that we have assumed simplistically that a solvent is either
interacting strongly enough to reorient with the BBD chromophore, or not. This
assumption can be re-expressed in terms of the average lifetime of the solvent-
solute interactions, where we assume that the 1-propanol-BBD interaction
persists for a time interval that is longer, on average, than the anisotropy decay
time of the reorienting moiety. If this is not the case, then we would expect in this
model to recover solvent-related “additional volume” that would be a fraction of
whole solvent molecule(s), and indeed, if we were to invoke this parameter, we
could achieve better correspondence between experiment and model. We do
not have any experimental information, however, on the lifetime of the 1-
propanol-BBD interaction, so we do not feel justified in pushing the model this
far.

It is not surprising, in our view, to find that there exist strong solvent-solute

interactions between BBD and 1-propanol. What is interesting is that our data

36

 

 

 

 

 

 

 

 

 

 

 

 

Wt % V (calculated n V 103 ton
Propanol from GXP- (Veeo+nVPror-l) calculated experimental

data) (ps) (081
100 388 3 425 200 209 :I: 3
88 321 2 355 191 201 :1: 1
76 271 1 285 171 190 :I: 1
65 227 0 215 137 171 i 2
55 214 0 215 139 166:1:1
45 208 0 215 135 156 :I:1
35 236 1 285 164 163 :1: 4
26 272 1 285 144 164 :1: 1
17 270 1 285 117 132:1:1

8 261 1 285 87 92 :1: 1

 

 

 

 

 

 

Table 2.1 - Solvent association calculations for V = meb. + nV.o.v.m.

Values of V calculated from experimental ton times are shown In the
second column, and from these data we can estimate the number of 1-
propanol molecules that reorient with the BBD chromophore. From this
estimate of n and experimental viscosity data (from Ref 52"5"), we calculate a

model T03.

37

 

 

are consistent with the extent of solvent-solute interactions varying in a non-
monotonic manner with solvent system composition. This finding implies that,
over a significant composition range for the 1-propanol/water system, the
interactions between water and 1-propanol are sufficiently strong to preclude 1-
propanol interacting strongly with the BBD chromophore. We note that the
formation of an azeotrope for a binary solvent system implies significant
interactions between the two solvents, and for 1-propanol and water, the
azeotrope-forming composition range lies between 28 to 70 weight percent 1-
propanol. In this solvent system composition range, it is possible that the most
stable solvent configuration(s) are those for which 1-propanol and water are
interacting more strongly with themselves than with BBD. In concentration
regions away from where an azeotrope forms, the individual solvent molecules
could, in principle, interact more strongly with BBD.

The above explanation provides qualitative agreement with our
experimental data, but it cannot provide more detail, based in part on the dearth
of molecular-scale information extant on the nature of azeotrope(s) between 1-
propanol and water. It appears that the strength of interactions between BBD
and the 1-propanol/water system are closely matched with 1-propanol-water
interactions, and that in all cases these interactions must persist over a time
scale that is consistent with hydrogen bond formation and breakage. Regardless
of the inability to provide a detailed picture of the solvation dynamics of BBD in
the 1-propan0l/water system, it is clear that depending on the amount of alcohol

present in solution, there can be significant intermolecular interactions between

38

 

chromophores and solvents, and these interactions can, in principle, have some

influence on the formation of micelles.

39

Conclusions

We have examined the fluorescence lifetime and rotational diffusion
dynamics of BBD in the 1-propanoVwater solvent system. The increase of the
fluorescence lifetime of BBD with increasing 1-propanol concentration is fully
consistent with the known behavior of chromophores containing an NBD group,
and indicates that the persistence time of any local organization is less than
nanoseconds. Modeling the reorientation behavior of BBD in the context of the
modified DSE model indicates that, for higher concentrations of 1-propanol, the
experimental anisotropy decay times are significantly longer than expected.
Because of the non-monotonic nature of the reorientation data, we understand
this anomalous behavior in the context of strong, associative solvent-solute
interactions, with the persistence time of the interactions being on the order of
the measured reorientation time. While strong solvent-solute association has

5'25”, this is a particularly pronounced example of solvent-

been seen before
composition dependent interactions, and we note that the solvent-solute
interactions are weakest in the 1-propanol concentration regime where
azeotropes are formed with water. It is clear that in the 1-propanoVwater solvent
system there is discernible microscopic solvent heterogeneity, with the details of

this transient solution phase structure depending sensitively on the composition

of the solvent system.

40

Literature Cited

(1) DelaCruz, J. L.; Blanchard, G. J. Journal of Physical Chemistry B
2003, 107, 7102.

(2) Kelepouris, L.; Blanchard, G. J. Journal of Physical Chemistry B
2002, 106, 6600.

(3) Kelepouris, L.; Blanchard, G. J. Journal of Physical Chemistry B
2003, 107,1079.

(4) Stevenson, S. A.; Blanchard, G. J. In Preparation.

(5) Dela Cruz, J. L.; Blanchard, G. J. Journal of Physical Chemistry A
2001, 105, 9328.

q a;

(6) Goldie, S. N.; Blanchard, G. J. Joumal of Physical Chemistry A
2001, 105, 6785.

(7) Goldie, S. N.; Blanchard, G. J. Journal of Physical ChemistryA
1999, 103, 999.

(8) Jiang, Y.; Blanchard, G. J. Journal of Physical Chemistry1994, 98,
941 1 .

(9) Jiang, Y.; Blanchard, G. J. Journal of Physical Chemistry 1995, 99,
7904.

(10) Heilweil, E. J.; Casassa, M. P.; Cavanagh, R. R.; Stephenson, J. C.
Annual Review of Physical Chemistry 1989, 40, 143.

(11) Heilweil, E. J.; Cavanagh, R. R.; Stephenson, J. C. Chemical
Physics Letters 1987, 134, 181.

(12) Casassa, M. P.; Heilweil, E. J.; Stephenson, J. C.; Cavanagh, R. R.
Journal of Chemical Physics 1986, 84, 2361.

(13) Cavanagh, R. R.; Heilweil, E. J.; Stephenson, J. C. Journal of
Electron Spectroscopy and Related Phenomena 1987, 45, 31.

(14) McCarthy, P. K.; Blanchard, G. J. Journal of Physical Chemistry
1995, 99, 17748.

(15) McCarthy, P. K.; Blanchard, G. J. Journal of Physical Chemistry
1996, 100, 5182.

41

(16) McCarthy, P. K.; Blanchard, G. J. Joumal of Physical Chemistry
1996, 100. 14592.

(17) Millar, D. P.; Shah, R.; Zewail, A. H. Chemical Physics Letters
1979, 66, 435.

(18) Goldie, S. N.; Blanchard, G. J. Book of Abstracts, 213th ACS
National Meeting, San Francisco, April 13-17 1997, ANYL.

(19) Fleming, G. R.; Morris, J. M.; Robinson, G. W. Chemical Physics
1976, 17,91.

(20) Pauls, S. W.; Hedstrom, J. F.; Johnson, C. K. Chemical Physics
1998, 237, 205.

(21) Truong, T. V.; Shen, Y. R. Journal of Chemical Physics 2005, 122,
091104/1.

(22) Balabai, N.; Sukharevsky, A.; Read, l.; Strazisar, B.; Kumikova, M.;
Hartman, R. S.; Coalson, R. D; Waldeck, D. H. Journal of Molecular
Liquids 1998, 77,37.

(23) Hartman, R. S.; Konitsky, W. M.; Waldeck, D. H. Journal of the
American Chemical Society1993, 115, 9692.

(24) Sanders, M. J.; Wirth, M. J. Chemical Physics Letters 1983, 101,
361.

(25) Mukherjee, A.; Bagchi, B. PhysChemComm 2003, 28.

(26) Von Jena, A.; Lessing, H. E. Chemical Physics Letters 1981, 78,
187.

(27) Dela Cruz, J. L.; Blanchard, G. J. Journal of Physical Chemistry A
2002, 106, 10718.

(28) Srinivas, G.; Bhattacharyya, S.; Bagchi, B. Journal of Chemical
Physics 1999, 110, 4477.

(29) Labhart, H.; Pantke, E. R. Chemical Physics Letters 1973, 23, 482.
(30) Blanchard, G. J. Journal of Chemical Physics 1987, 87, 6802.

(31) Blanchard, G. J. Journal of Physical Chemistry 1988, 92, 6303.
(32) Blanchard, G. J. Analytical Chemistry 1989, 61, 2394.

42

(33) Blanchard, G. J.; Wirth, M. J. Journal of Physical Chemistry 1986,
90, 2521.

(34) Templeton, E. F. G.; Quitevis, E. L.; Kenney-Wallace, G. A. Journal
of Physical Chemistry 1985, 89, 3238.

(35) Chuang, T. J.; Eisenthal, K. B. Journal of Chemical Physics 1972,
57, 5094.

(36) Debye, P. Polar Molecules. In Chemical Catalog Go. New York,
1929.

(37) Perrin, F. Journal de Physique et la Radium 1936, 7, 1.

(38) Hu, C.-M.; Zwanzig, R. Journal of Chemical Physics 1974, 60,
4354.

(39) Zwanzig, R.; Harrison, A. K. Journal of Chemical Physics 1985, 83,
5861 .

(40) Fery-Forgues, 8.; Fayet, J.-P.; Lopez, A. Joumal of Photochemistry
and Photobiology, A: Chemistry 1993, 70, 229.

(41 ) Lin, 8.; Struve, W. S. Photochemistry and Photobiology 1991 , 54,
361 .

(42) Mukherjee, S.; Chattopadhyay, A. Journal of Fluorescence 1995, 5,
237.

(43) Mukherjee, S.; Chattopadhyay, A.; Samanta, A.; Soujanya, T.
Journal of Physical Chemistry 1994, 98, 2809.

(44) DeWitt, L.; Blanchard, G. J.; LeGoff, E.; Benz, M. E.; Liao, J. H.;
Kanatzidis, M. G. Journal of the American Chemical Society 1993, 1 15,
12158.

(45) Froimowitz, M. Bio Techniques FIELD Publication Date:1993, 14,
1010. .

(46) Edward, J. T. Journal of Chemical Education 1970, 47, 261.

(47) Bhattacharyya, S.; Bagchi, B. Journal of Chemical Physics 2001,
115, 9065.

(48) Dutt, G. B.; Doraiswamy; Periasamy, N. Journal of Chemical
Physics 1991, 94, 5360.

43

(49) Alavi, D. S.; Waldeck, D. H. Journal of Physical Chemistry 1 991 ,
95, 4848.

(50) Mukherjee, A.; Bagchi, B. Journal of Physical Chemistry B 2001 ,
105, 9581 .

(51) Beddard, G. 8.; Doust, T.; Hudales, J. Nature (London, United
Kingdom) 1981, 294, 145.

(52) CRC Handbook of Chemistry and Physics, 1980-1981.

(53) Mikhail, S. 2.; Kimel, W. R. Journal of Chemical and Engineering
Data 1963, 8, 323.

(54) Wensink, E. J. W.; Hoffmann, A. C.; van Maaren, P. J.; van der
Spoel, D. Journal of Chemical Physics 2003, 1 19, 7308.

44

 

Chapter 3

INVESTIGATING THE MlCELLE-SOLUTION INTERFACIAL REGION USING
FLUORESCENCE RESONANCE ENERGY TRANSFER

Introduction

Surfactants have attracted considerable interest for a number of years due
to their many uses, ranging from chemical separations to industrial and home-
use detergents. Among the interesting properties of surfactants is their ability to
form micelles of various shapes, depending on the specific surfactant and its
concentration. Despite the substantial micelle literature extant, a detailed
understanding of the structure and dynamics of this family of systems remains to
be achieved. We are interested in understanding the intermediate region
between micelle head groups and the bulk solvent surrounding them because it
is this interface that mediates exchange processes and interactions between the
micelles and other species in solution. Depending on the strength of such
interactions, it is also possible that the interior of the micelle will be affected, and
this is an issue we address experimentally. Because there is no direct means to
probe the micelle-solution interface electrochemically, we are interested in
determining the utility of fluorescence spectroscopy for that purpose.

Fluorescence spectroscopy finds wide use because of its characteristically
high sensitivity combined with the range of chemical properties available with the
fluorophores used. For measurements in heterogeneous systems, it is possible
to use fluorescent compounds that will partition selectively into a given phase, or

will reside at the interface between two phases. This capability, when combined

45

with fluorescence resonance energy transfer (FRET) measurements, has been
used in the study of heterogeneous systems such as micelles, vesicles, and lipid
bilayers. "23 The topology of these systems can significantly influence the
efficiency of energy transfer since the dipolar coupling process that mediates
energy transfer depends on both the average distance and orientation of the
molecules involved. Energy transfer measurements can be used as a
“spectroscopic ruler" to estimate average distances between optical donor and
acceptor, and thus we can gain information on the extent to which micelle and
acceptor interact. Time-domain reorientation measurements can provide
information on the microenvironments that exist within a heterogeneous system.
Micellar systems are ideal candidates for energy transfer experiments, and
previous work in this area has examined energy transfer between two molecules
sequestered inside a micelle, between a molecule inside the micelle and a
molecule that is part of the surfactant, and between two molecules on the surface
of the micelle. ‘2'“:‘9'2‘ Most of these studies have focused on energy transfer
between molecules in the same region of the micelle. We are particularly
interested in the interfacial region of micelles, and understanding the
intermolecular dynamics in micellar systems will hopefully shed some light on
more complex biomimetic systems.

We have sequestered perylene inside sodium dodecyl sulfate (SDS,
anionic) and cetyltrimethylammonium bromide (CTAB, cationic) micelles, and
have used the steady state and time-resolved fluorescence of this chromophore

to obtain information on how different acceptor chromophores in the aqueous

46

 

be
38
di:
SP

OP

aCl
chl
inle
inle
the
a lL

of ti

 

 

phase interact with the donor. Perylene is a hydrophobic chromophore that has
been studied extensively, making it an obvious choice for a probe to be
sequestered within micelles. The two ionic acceptor chromophores, anionic
disodium fluorescein (DSF) and cationic rhodamine 110 (R110), have
spectroscopic properties well suited to their role as optical acceptors, and their
opposite ionic charges make them useful for studying interactions with micelles
formed by the two ionic surfactants we use. By varying the concentration of the
acceptor chromophores, and the ionic charges present on both the acceptor
chromophores and the micelle head groups, we can evaluate the role of ionic
interactions in mediating molecular communication across the micelle-solution
interface. From these data we can estimate the association constant between
the chromophores and the micelles. Reorientation measurements of perylene as
a function of optical acceptor concentration point to the role of the palisades layer

of the micelles in mediating the properties of the micelle interior regions.

47

Experimental

Materials. The surfactants sodium dodecyl sulfate (SDS) and
cetyltrimethylammonium bromide (CTAB) were obtained from Sigma-Aldrich, Inc.
and used as received. Perylene (Aldrich), disodium fluorescein (Aldrich) and
rhodamine 110 chloride (Exciton) were checked for fluorescent impurities and
used as received. The structures of the chromophores used in this chapter can
be seen in Figure 3.1. The perylene concentration for all solutions was 10 pM.
The micellar solution temperatures were maintained at 298 :0.5 K (Neslab
EX100-DD) for all time-resolved fluorescence measurements. For the SDS
solutions, perylene was dissolved in a 50/50 solution of butanol and methanol
(3x10‘5 M perylene concentration) before addition of surfactant and, similarly, for
CTAB solutions the perylene was dissolved in acetonitrile (3x10'5 M) before
addition of the surfactant. These cosolvents were required to get perylene into
the micellar solutions. For the formation of micelles, we used 17 mM SDS, which
is twice the critical micelle concentration, and 2 mM CTAB, also corresponding to
twice the critical micelle concentration. Cryo-TEM images (vide infra) show
uniform size and shape of resulting spherical micelles in solution. The ionic
strengths of the solutions varied with composition.
Time-Correlated Single Photon Counting Measurements. Fluorescence lifetime
and anisotropy measurements of perylene sequestered within SDS and CTAB
micelles were made using a time-correlated single photon counting (T CSPC)
instrument that has been described in detail previously.25 The excitation

wavelength was 420 nm and the emission was collected at 470 nm.

48

C C ‘ coon
C O ‘O
HZN 0 NH;

perylene Disodium fluorescein Rhodamine 110
DSF R1 10

 

 

O\SO3' Na+
Sodium dodecyl sulfate (SDS)

W\/\/\/\/\/\/N+(CH3)3 B"
Cetyl trimethylammonium bromide (CTAB)

Figure 3.1 - Structures of the chromophores used in thls chapter.

49

Steady-State Spectroscopy. Absorption spectra were recorded using a
Cary model 300 double-beam UV-visible absorption spectrometer, with 1 nm
spectral resolution. Spontaneous emission spectra were acquired with a Spex
Fluorolog 3 spectrometer, with both excitation and emission monochromators set
to 3 nm resolution.
Cryo-TEM imaging. Cryogenic TEM images were acquired on our samples by
Dow-Corning Analytical Solutions, Auburn Hills, MI. To prepare the samples, ca.
2.3 mL of the micelle-containing solution was loaded on a holey carbon film
coated Cu TEM grid using a micropipette. The excess fluid on the grid surface
was removed by blotting the surface with a filter paper make an aqueous thin film
for the TEM. The grid was immersed in liquid ethane contained in a small vessel
that itself was housed in a larger liquid nitrogen vessel. The water film on the
grid was vitrified to avoid water crystallization. The sample was loaded into the
TEM (JEOL JEM 2100F) and the morphology was observed below 104 K. The
digital images were acquired using a Gatan CCD camera attached at the bottom
of the TEM column and Digital Micrograph software. The cryo-TEM images are

shown in Figure 3.2.

50

 

showing a

Figure 3.2 - Cryo-TEM Images of SDS (a) and CTAB (b) micelles,

range of sizes less than 100nm diameter.

51

Results and Discussion

The objective of this chapter is to better understand the interfacial region
between the micelle head groups and the bulk solution. We are concerned with
the role of ionic charge in determining the interactions between ionic micelle
head groups and constituents in the bulk solution. We probe this region by
monitoring the effect that ionic optical acceptor molecules in the aqueous phase
have on a fluorophore sequestered within the micelle. Excitation transport
between the donor and the solution phase acceptor chromophore provides
insight into the extent of interaction between acceptor and micelle, and the
motional properties of the chromophore sequestered within the micelle relate to
micellar organization and local viscosity. We detail below how our results point
to discernible heterogeneity in the micellar solutions and changes in the nonpolar
region of the micelle with the addition of an optical acceptor chromophore.

Before discussing our experimental data in detail, a word is in order on
what is known about the micelle/solution interface. While it may be tempting to
consider the interface between micelles and the bulk solution to be relatively well
defined, several issues serve to complicate this picture. The first is surfactant
exchange between the solution phase and the micelle. The typical residence
time for a surfactant molecule in the micelle assembly will, of course, vary
depending on the specific system, but such times are on the order of
nanoseconds to microseconds. 2° These interfaces are also not well represented

as a single, charged “ball”. If this were the case, head group ionic repulsions

52

would preclude micelle formation. Based on a large body of literature and
previous work in the Blanchard group,“'27 it is clear that the counter ions in
solution play a significant screening role and mediate ionic interactions between
surfactant head groups. The interface between the micelle and the bulk solution
is not abrupt, progressing from a non-polar micelle interior, through an ionically
compensated region, through a solvent “layer" that likely exhibits organization
analogous to that of the electric double layer, and finally to bulk solution. This
intermediate region is termed the “palisades layer”, 26'2”" and it is the details of
this intermediate region that are of primary interest to us in this work.
Fluorescence lifetime measurements. Our experiments are designed to
understand whether or not the ionic charge of micelle head groups serves as a
point of strong interaction with ionic constituents in solution. To make this
determination, we use fluorescence lifetime measurements to characterize
excitation transport dynamics in micelle-containing solutions. The concentration
of optical donor in the micelles is such that we estimate there to be a single
perylene chromophore in approximately 1% of the micelles. The optical acceptor
concentrations we use are such that in a homogeneous solution, the average
distance between donor and acceptor would be 150 A or more. Under these
experimental conditions, very little donor-acceptor excitation transfer will be seen,
and we will observe acceptor concentration-dependent changes in the donor
fluorescence lifetime only if the optical acceptor interacts preferentially with the

micelles.

53

The transfer of energy between molecules can occur by a number of
mechanisms, ranging from collisional quenching to dipolar coupling. We are
concerned in this work with the latter phenomenon because of the characteristic
length scale over which it operates. The transfer of energy by dipolar coupling

has been described by Fcrster, 3‘

._ 6
k 12"? -.°88x10"j A ("2’0 f(v)dl7="2kp[R°] 3.1

 

DA: n"R6 n4 T?—
where km is the rate constant for the transfer of energy between donor and
acceptor. The K term is a function of the relative orientation of the donor and
acceptor transition dipole moments, and for our experimental conditions, <K2>=
2/3. The term k,” is the donor radiative decay rate constant in the absence of
optical acceptor, n is the refractive index and R is the intermolecular D-A
distance. The integral term is a measure of the spectral overlap between the
emission spectrum of the optical donor and the absorption spectrum of the
acceptor. The spectral overlap term, combined with the several constants on the
right hand side of Eq. 3.1 is typically referred to as the critical radius, R0, a
quantity which serves as an effective gauge of the operating distance of this
dipolar coupling process. While the term R0 will vary for each donor-acceptor
pair, for most organic chromophore pairs, R0 ~ 50 A. ”33'3"”
We have used the optical donor perylene and two different acceptor
molecules, DSF (anionic) and R110 (cationic). The acceptor molecules were

chosen because of their ionic charge and their absorption spectral profiles, which

overlap significantly with the emission spectrum of perylene (Figure 3.3). Based

54

Fig

 

DSF absorption

. R110 absorption
P \

/

  
  
   
   

perylene emission

- perylene absorption

extinction coefficient (Umol-cm)

 

  

 

 

350 375 400 425 450 475 500 525 550 575

wavelength (nm)

Figure 3.3 - Normalized absorption and spontaneous emission spectra of
perylene, and absorption spectra of DSF and R110.

55

on the spectral overlap, and the extinction and fluorescence quantum yield data
on these molecules, we estimate R0 ~ 54 A for the perylene/DSF pair and R0 ~
51 A for the perylene/R110 pair. We excite perylene at 420 nm and measure its
fluorescence lifetime at ca. 470 nm, without interference from acceptor emission.
The measurement of donor lifetime as a function of acceptor concentration is a
direct means of evaluating variations in km, since this rate constant will operate
in an additive manner with the intramolecular decay rate constant(s) that govern
the depopulation of the donor excited state. Since the concentration of perylene
is held constant at 10'5 M, the amount of energy transfer between two donor
molecules should be negligible because of the small fraction of micelles that
contain a perylene molecule. Even if this process were significant, it would
appear as a depolarization of the perylene fluorescence, but not a diminution of
its lifetime. 33

We consider first the interactions between (anionic) DSF in solution and
perylene in (anionic) SDS micelles. As shown in Table 3.1, the fluorescence
lifetime of perylene decreases with increasing DSF solution concentration. This
is an expected result. In addition to the decrease in fluorescence lifetime, the
increasing concentration also affects the functionality of the perylene decay. For
10’3 M DSF, we observe a two-component perylene fluorescence lifetime. We
understand this behavior in the context of interactions between the acceptor
chromophore and the micelle, and will return to a discussion of this point after we

consider the analogous data for other systems.

56

 

 

 

 

 

 

 

 

 

Acceptor Concentration rm (ps) 142 (ps)
- - 5696 e 1 --

DSF 1? 5491 :1; 1
DSF 10* 5278 :1: 1
DSF 10'3 888 :1: 10 4362 e 3
R110 10'5 5468 :I: 2
11110 10“ 5197412
R1 10 10'3 953 :1; 4 4749: 4

 

 

 

 

 

 

Table 3.1 - Fluorescence lifetime data for perylene sequestered In SDS
micelles as a function of optical acceptor concentration. The data reported
here are the averages of 6 Independent determinations, and the
uncertainties reported are :1: 1o.

57

 

We compare these data to those for perylene in SDS micelles and (cationic)
R110 in solution. These results are qualitatively the same as for the
perylene/SDS/DSF system, including the onset of a two-component perylene
fluorescence lifetime for 10'3 M R110 acceptor. If simple ionic interactions were
responsible for these data, we would expect a substantial difference the perylene
lifetime with these two acceptors. The similarity of the perylene lifetimes in the
presence of either acceptor points to the significant role that counter ions must
play in mediating interactions between charged species at the micelle surface.
We have also studied perylene in (cationic) CTAB micelles, in the
presence of the two acceptor chromophores. For the solutions containing DSF,
the perylene lifetime exhibits qualitatively the same acceptor concentration-
dependence as is seen in SDS. For acceptor concentrations less than 10'3 M we
observe the expected acceptor concentration-dependent single exponential
decay, and for 10’3 M acceptor we observe a two-component decay. For R110 in
solution and perylene in CTAB, we recover essentially the same results as for the
other systems, save for 10‘3 M R110, where two different double exponential
lifetimes are observed (Table 3.2). The fact that two different decay
functionalities can be obtained reproducibly and interchangeably suggest that we
are near some phase transition for this system. In both cases a double
exponential is observed, consistent with two different environments for the
perylene, but the lifetime data in and of themselves are not sufficient to provide
any structural detail on these environments. We consider the result for 10" M

R110 with perylene in CTAB to be a variant of the results we have obtained for

58

 

 

 

 

 

 

 

 

 

 

Acceptor Concentration rm (ps) 142 (ps)
- - 5529 :I: 1
DSF 10'5 5374 :I: 1
DSF 10‘r 5096 a: 1
DSF 10'3 701 :1: 4 3877 :1: 3
R110 10'5 5470 :l:1
R110 10" 5243 :I: 1
R110 10’3 1315:25 3936::7
2040211 5827116

 

 

 

 

 

 

Table 3.2. Fluorescence lifetime data for perylene sequestered In CTAB
micelles as a function of optical acceptor concentration. The data reported
here are the averages of 4 independent determinations, and the
uncertalntles reported are :l: to.

59

the other systems. Overall, the similarity of these excitation transport data for
cationic and anionic micelles, and cationic and anionic acceptors points to the
significant screening role that counter-ions must play in determining the
properties of the palisades region of the micelle/solvent interface.

In addition to qualitative insight into excitation transport between the
interior and exterior region of the micelles, these lifetime data can also provide
quantitative insight into the interfacial region. We consider that acceptors form a
complex with the micelle. In this picture, the acceptors are either in very close
spatial proximity to the donor when complexed, or too far distant to transfer
energy efficiently when free. This is, of course, an oversimplification, but it allows
for the ready use of the Fcrster equation in the estimation of association
constants for these systems.

It is first important to consider the approximate stoichiometry of the donor,
acceptor and micelles in our solutions. As noted above, the perylene donor
concentration is such that there is one donor per ~100 micelles. As we increase
the acceptor concentration from 10'5 M to 10'3 M, the acceptor-to-micelle
stoichiometry changes from one acceptor per ~ 100 micelles to ~ 4 acceptors per
micelle for SDS. We base this estimate on the micelle concentration being

[micelle] ~ ” CMC , 3.2
surfactant molecules per micelle

 

Where n is the number of CMC concentrations we use (n=2 here) for SDS (CMC

= 8.5 mM), there are ~63 surfactant molecules per micelle, and for CTAB (CMC =

60

0.8 mM) there are ~90 molecules per micelle.3‘"35 This estimate yields [micelle]~

0.27 mM for SDS and 0.02 mM for CTAB.

Because of the perylenezmicelle ratio of 1:100, for a 10'5 M acceptor
concentration, there is one acceptor per ~100 micelles, so only 0.01% of the
micelles will have both a donor sequestered within and an acceptor complexed to
them. Under these conditions, there will be essentially no donor-containing
micelles that are complexed to more than one acceptor. We can thus
approximate safely that for low acceptor concentrations, the equilibrium process
we monitor with our donor lifetime measurements corresponds to interactions
between single donors and single acceptors. For 10'3 M acceptor
concentrations, the situation is fundamentally different. There are ~4 acceptor
molecules per micelle, making the association between micelles and multiple
acceptors the dominant interaction, and this situation is manifested by the
change in the donor decay functionality from single exponential to double
exponenfiaL

Before considering the information content of these data, we need to
evaluate the sizes of the micelles under investigation. The cryo—TEM data on our
micelles (Figure 3.2) indicate that the micelles are in the sub-10 nm range, but
we are not able to extract precise average diameters. It is important to consider
that these micelles have experienced vitrification, so it is not unreasonable to
expect that there has been some structural perturbation prior to acquisition of the
images. The larger point of these images is that we do indeed have nanoscale

micelles present in our solutions, and with their existence established, we will use

61

 

the more precise estimates of micelle size that have been reported elsewhere.
Specifically, the radius of SDS is taken to be 19 A and the radius of CTAB is
taken to be 25 A for the purposes of the calculations we present below.”37

We assert that the lifetimes that we measure are actually weighted
averages of the fluorescence lifetime of perylene in micelles complexed with one
acceptor and the lifetime of perylene in an uncomplexed micelle. We take the
experimentally determined fluorescence lifetime as the weighted average of the
free form 1.... = (k.")’1 and the complexed form, where rcpt. = (k.D + km)", with k0,.

being calculated for R =21 A (SDS) or R =25 A (CTAB) and R; as given above.

Tabs = nfreeTfree + ncpIxTcpIx
n = [M] ,, _ lMAl 3-3

1... [M] + [MAI’ cp'x "' “—lMl + lMAl
From these calculations, we recover the fractional contributions from the
free and complexed forms of the perylene-containing micelles. For the acceptor

association with the micelles, the relevant equilibrium process is:

= [MA] = ncplx
[M ][A] n [A]

 

M+A K MA K (4)

 

 

ree

We know [M] and [A] from solution composition, and determine the ratio of
[MA]/[M] experimentally from the lifetime measurements. We determine the
quantities rim and new (Eq. 3.3), and use them to calculate K. We estimate the
equilibrium constants for the donor/acceptor systems in both of the micellar
solutions (Table 3.3). We find that for either acceptor present at 10'5 M in SDS
micelles, K ~ 2000 M", and in CTAB micelles, K ~ 1300 M" under these same

conditions.

62

 

 

 

 

 

 

 

 

 

 

Micelle Acceptor Concentration Kaq rim nap),
SDS DSF 10-5 1833 0.982 0.018
SDS DSF 10" 395 0.962 0.038
SDS R1 1 O 1 0‘5 2040 0.980 0.020
SDS R1 10 10" 482 0.954 0.046

CTAB DSF 10'5 1420 0.986 0.014

CTAB DSF 1 0““ 428 0.959 0.041

CTAB R110 10'r 1215 0.988 0.012

CTAB R1 10 1 04 277 0.973 0.027

 

 

 

 

 

 

Table 3.3 - Equilibrium constant calculations for DSF and R110 In SDS and

CTAB.

63

 

If this same treatment is applied to solutions containing 10“ M DSF and R110,
we obtain values of K in the range of 250 M" to 500 M", depending on the
system. We do not believe the formation constant(s) for the 10" M acceptor
solutions to be different than those for the 10’5 M solutions. Rather, we view the
value of K ~ 250 — 500 M" as the overall equilibrium constant for multiple

equilibrium processes. For a multiple equilibrium process,

 

M+A£MA+A—Kz—*MAZ K= K,K2 (5)

From the experiments with 10'5 M acceptor, K1 ~ 2000, and for the 10" M
acceptor solutions, K1K2 ~ 400, leading to a value of K2 ~ 0.2, its exact value
depending on the system. While it is normal for the equilibrium constants to
diminish monotonically with the addition of complexing species, the difference
between K1 and K2 for acceptor/micelle complexation is substantial. It is possible
that the size of the acceptors and facile translational motion of the acceptor on
the micelle “surface” give rise to a significant contribution from steric effects for
the association of multiple acceptors with a given micelle.

The Fayer group’s elegant work on excitation transport in micellar media
has shown that a Green’s function describes the donor population decay

9'24'32'33'38'39 It is fair to question whether the data we report here are

dynamics.
characterized better by a Green’s function or a simple sum of exponentials. We
have attempted to fit our data to the appropriate Green’s function, and find that a
sum of exponentials accounts most accurately for the decay functionality of our

systems. This finding indicates that the observed decay is most likely the result

of two different perylene populations, i.e. those where one acceptor is associated

64

with the micelle, and one where two acceptors are associated with the micelle.
This finding is consistent with multiple micelle-acceptor association equilibria
contributing to our experimental findings at high acceptor concentrations. We
base this assertion on the comparatively abrupt change in functionality between
one and two component decays. To evaluate the possibilities of either a phase
transition for these micelles at high acceptor concentrations, or potential changes
in organization within the micelles, we have investigated the motional freedom of
the perylene molecule sequestered within the micelles as a function of acceptor
concentration.

Despite any subtle differences in detail, it is important to note the overall
similarity of the results for both micelles, and for both anionic and cationic
chromophores. These data demonstrate clearly that it is not direct ionic
interactions that dominate in these systems. Rather, the mediating effect that the
solution phase counterions play in the interactions between micelles and other
moieties in solution dominates the observed behavior.

Rotational diffusion within the micelles. To understand the influence of the
acceptor molecules on the motional freedom of the sequestered donor, we have
studied the rotational diffusion dynamics of perylene within the micelles as a
function of acceptor concentration and micelle identity. Rotational diffusion has
been used extensively to interrogate local organization in solutions, because of
the well developed theoretical framework for interpretation of the data and the
relatively high information content of the experimental signals. ‘°'“ Our data

indicate that the two acceptor molecules influence donor motion differently.

65

We obtain fluorescence depolarization data by exciting perylene with a
vertically polarized light pulse and collecting the emission decay transients
polarized parallel and perpendicular to the excitation polarization. These data

are combined to produce the induced orientational anisotropy function, R(t),

_ ’(t) " It“)
H“) ' litl+2tltl

 

(2)

where l"(t) and I i(t) are the polarized emission intensities. The chemical
information contained in these data lies in the decay time constant(s) of R(t), and
its zero-time value.

Before discussing these data further, we note that donor-donor excitation
transport can give rise to fluorescence depolarization”, the same physical
phenomenon that we use to characterize the molecular motion of perylene
sequestered within micelles. The DA excitation transport data (vide supra) point
to modest donor-acceptor excitation transport, and given the low perylene
concentrations used (~1 perylene per 100 micelles), we believe that fluorescence
depolarization resulting from donor-donor transport does not contribute
measurably to our data.

Because we are studying heterogeneous systems, it is fair to consider
which model is most appropriate for the treatment of the anisotropy data. For
chromophores tethered to a substantially larger entity, such as a protein, or
imbedded in a mono- or bilayer structure, the hindered rotor model is
appropriate.MS In this model the chromophore is bound at one point within the
system and its motion is considered in the context of the conic volume it sweeps

out. For the micellar systems we are concerned with here, the hindered rotor

66

model is not appropriate because the chromophore is not bound to a specific
point in or on the micelle. Rather, perylene is free to diffuse within the micelle,
and we treat this system in the context of perylene reorienting in a quasi-bulk
solvent. The model most appropriate for this type of system is the modified
Debye-Stokes-Einstein (DSE) equation, ‘6 which is described in more detail in
chapter 1 (equation 1.3).

For perylene the hydrodynamic volume is 225 A3,“7 T = 300 K, S = 0.69 (prolate
rotor based on single exponential decay). “"9 We note that our experimental
anisotropy data are all single exponential decays, allowing us to conclude that
perylene reorients as a prolate rotor in the micelles we have studied. Perylene
has been shown previously to reorient either as a prolate rotor (single
exponential anisotropy decay) or as an oblate rotor (double exponential
anisotropy decay), depending on its environment. ‘9'“ While it may seem
plausible to invoke a slip boundary condition for our experimental conditions,
experimental data for perylene in n-alkanes are consistent with behavior
intermediate between the stick and slip limits. From the modified DSE model, in
the stick limit we calculate the quantity tor/Tl = 78 ps/cP, and in the slip limit we
calculate 1an = 5.8 ps/cP. Experimentally we know from earlier work on
perylene in n—alkanes longer than C3, tor/11 = 12.7 a: 1.5 ps/cP, intermediate
between the stick and slip limit, and we use this experimental value to estimate

the viscosity of the micellar media.“"52 For perylene in SDS micelles, in the

67

 

 

 

 

 

 

 

 

 

 

 

 

Acceptor Concentration Ton 71 (CF)

- - 62:3 49:09
DSF 10'5 95 :I: 3 7.5 :t 1.3
DSF 10" 127 :I: 2 10.0 :I: 0.9
DSF 10'3 12814 10.1 :l:1.7
R110 103 123:1:5 9.75:1.6
R110 10“ 11613 91:15
R110 10'3 86:2 6.85:1.1

 

Table 3.4. Rotational diffusion time constants of perylene sequestered In
SDS mlcelles as a function of optical acceptor concentration. The data
reported here are the averages of 6 Independent determinations, and the
uncertainties reported are :1: 1o.

68

 

absence of acceptor chromophores in solution, we estimate the viscosity within
the micelle to be 11 = 4.9 :I: 0.9 CR and with the addition of anionic DSF acceptor,
the viscosity of the interior of the micelle varies monotonically between 7.5 cP
and 10.1 cP (Table 3.4). The viscosity of the aliphatic medium within the
micelles is substantially higher than the corresponding length n-alkane (~ 3.5 cP).
Some of this “excess” viscosity is likely due to the geometric constraints imposed
by the micellar structure itself. The increase in apparent viscosity with the
addition of acceptor suggests that the solution phase ionic species interact with
the micelle in such a way at to make the interior of the micelle less fluid. This
finding is consistent with either strong micelle-acceptor interactions mediating the
exchange of amphiphiles between solution and micelle, or a change in the lateral
mobility of the amphiphiles within the micellar structure. For the case of an
anionic acceptor (DSF) and anionic micelle head groups (SDS), we observe an
increase in the apparent micelle viscosity with the addition of acceptor, and a
plateau effect for 10“ and 10‘3 M acceptor concentrations. For the perylene in
SDS micelles, with the cationic R110 acceptor, we observe a different trend
(Table 3.4, Figure 3.4a). As the concentration of R110 is increased from 10’5 M
to 10'3 M, the micelle interior viscosity decreases from 9.7 cP to 6.8 cP. This
trend in micelle viscosity is opposite that seen for the anionic acceptor, pointing
to a structural role for charge-dependent interactions between the micelle head

groups and the solution phase acceptor.

69

 

 

 

 

 

 

 

 

 

Acceptor Concentration 103 TI (cP)

- - 100 2 5 7.9 2 1.5
DSF 10'5 50 2 1 3.9 2 0.7
DSF 10" 15323 12021.9
DSF 10’3 84 2 7 6.6 2 1.5
R110 10'5 14727 11.6222
R110 10" 11623 9021.6
R110 10’3 18022 14.2221

 

 

 

 

Table 3.5. Rotational diffusion time constants of perylene sequestered in
CTAB mlcelles as a function of optical acceptor concentration. The data
reported here are the averages of 4 independent determinations, and the
uncertainties reported are 2 1o.

70

 

For cationic micelles, we do not observe analogous monotonic trends with
increasing donor or acceptor concentration (Figure 3.4). For the native CTAB
micelle, in the absence of any optical acceptor, we recover a viscosity of 7.9 cP
for the micelle interior (Table 3.4). As anionic DSF acceptor is added to the
micelle-containing solution, the micelle interior viscosity ranges from 3.9 CF to
12.0 CF, but this trend does not vary linearly with acceptor concentration. With
the addition of the cationic acceptor R110, the micelle interior viscosity varies
between 9.0 cP and 14.2 CF, but again, this trend is not monotonic with
increasing acceptor concentration (Table 3.5, Figure 3.4). In these experiments,
we observe significant variations in the apparent viscosity of the micellar
environment with changes in solution phase acceptor concentration. This
situation could be the result of a number of phenomena, but it is significant that
the trends we observe are not monotonic. If these data were accounted for in the
context of variations in a system property such as ionic strength, we would
expect monotonic trends in the reorientation and lifetime data. In fact, the
reorientation data do not correlate with the fluorescence lifetime data, and the
addition of the cationic R110 acceptor causes anomalous, non-monotonic
behavior in both cationic and anionic micelles. Given that the reorientation

dynamics of perylene within the micelles depend primarily on the viscosity of the

71

 

 

 

 

6000 - a
E
l- a a
5000 -
B
' e
A 4000 F
is; .
c3000 -
P .
2000 F
1000 - 6
| [II I 4 l l I l I I I 1 l l
-5.5 -5.0 -4.5 -4.0 -3.5 -3.0
log1 0[acceptor]
6000
r. , . b
5000 , 5
4000 - e
@3000 -
P 2000 - a
1000 - a
_ O
O —'|_-/ll l I j l l I l l l l I
-5.5 -5.0 -4.5 -4.0 -3.5 -3.0

log 1 0[acceptor]

Figure 3.4 - Fluorescence lifetimes of perylene sequestered in mlcelles as a
function of optcal acceptor concentration. (a) SDS mlcelles, e = DSF, =

R110. (b) CTAB mlcelles, o = DSF,

= R110.

72

 

 

 

 

175 - a
150 -
s l i
3 125 . § 5
PS 100 - i
' El
75 -
- a
50 -
_|__// I . l . I . I . I 1 l .
-5.5 -5.0 -4.5 -4.0 -3.5 -3.0
log1 0[acceptor]
175 - '3 b
150 - § 5
’5? 125 -
3 . o
P#5 100 - 5
75 .. t
50 - o
_|_/,L l l . l . l . I . l .
~5.5 -5.0 -4.5 -4.0 -3.5 -3.0
log 10[acceptor]

Figure 3.5 - Rotational diffusion time constants of perylene sequestered in
mlcelles as a function of optcal acceptor concentration. (a) SDS micelles,

0 = DSF,

= R110. (b) CTAB micelles, o = DSF,

= R110.

73

micelle interior, our findings suggest a micelle structural rearrangement occurs
with the addition of R110. This rearrangement could either be structural, such as
a transformation from a spherical micelle to some other shape or attachment of
multiple R110 molecules to the micelle, or it could be compositional. We have
added a small amount of butanol cosolvent to the system, approximately 20% by
volume, to dissolve the perylene chromophore and thereby allow its incorporation
into the micelles. If the presence of butanol plays a role in the structure of the
micelle, the addition of acceptor molecules would alter the ionic strength of the
micelle-containing solution, thus changing the extent to which the butanol
cosolvent would partition into the micelles. Such an effect would scale with the
solution ionic strength, and thus a monotonic change in to, would be expected,
and that trend should be qualitatively the same for both acceptor chromophores.
The fact that the reorientation dynamics of perylene vary with the
presence of the two acceptors suggests an energetically significant interaction
between the acceptors and some constituent(s) of the micelles. This finding is
consistent with the substantial first association constants we measure for micelle-
acceptor interactions. The amphiphiles SDS and CTAB may undergo hydrogen
bonding with the solvent, but their ionic charge would likely cause them to be in
close association with their counter ions. The acceptor chromophores could, in
principle, participate in hydrogen bonding with butanol, and such an interaction
would be different for the two acceptors. Any interaction such as this, however,

should scale linearly with acceptor concentration (at least to the point of

74

saturation), and this expectation is counter to the experimental data we have for
R110. We are thus in need of acquiring further information to understand the
perylene/acceptor/CTAB data fully, but there is a substantial literature on the
modification of micelle properties with n-alkanols, the usual result being a
reduction in the viscosity of the micelle with increasing alkanol concentration.48 It
is possible that the presence of butanol in the solution and/or micelles gives rise
to structural variations akin to a phase transition within the micelle, and this is a
point which will require further investigation to resolve.

Regardless of the details of the interactions responsible for the anomalous
reorientation of perylene in CTAB micelles, we find in all cases that the observed
viscosity within the micelles is significantly higher than that of the corresponding
length alkane solvent. The microviscosities of micellar environments have been
investigated before, and our values are fully consistent with other literature
reports. “'55 The fact that the viscosity within micelles is higher than that seen
for the corresponding alkane, and more on a par with that seen for the
corresponding n-alkanol, underscores the role that the solution-head group
interactions in the palisades layer region play in mediating the behavior of the
hydrophobic micelle inner region. For bulk liquid phase alkanols, hydrogen
bonding between the hydroxyl groups functions in concert with van der Waals
interactions of the aliphatic tails to determine the viscosity of the medium. In the
case of micelles, the palisades layer must be sufficiently charge compensating to
prevent repulsive Coulombic forces from interfering with micelle formation. This

balance of ionic and H-bonding forces in the palisades layer will function in

75

concert with van der Waals forces within the micelle interior to yield a relatively
viscous environment. The alteration of the ionic strength of the solution will
change the balance of forces that operate within the palisades layer and,
depending on the details of the interactions, the micelle structure may be altered

as a result.

76

Conclusions

We have studied the fluorescence lifetime and reorientation dynamics of
perylene in the aqueous micellar systems SDS and CTAB in the presence of
varying concentrations of cationic (R110) and anionic (DSF) optical acceptor
molecules. The decrease in the lifetime of perylene in the presence of the
acceptors is fully consistent with dipolar excitation transport and from these data
we can infer the association constant(s) for micelle-acceptor interactions.
Comparing data for cationic and anionic micelles and both cationic and anionic
acceptors, we find that direct ionic interactions play little role in our data. In
homogeneous solutions, the average intermolecular distance is such that
excitation transport would be expected not to play a significant role, and our
observation of this effect in micellar solutions is an indication of interactions
between the charged optical acceptors and the palisades layer of the micelles.
The similarity of the behavior seen for all micelle/acceptor combinations points to
the important role that spectator ion charge compensation plays in micelle
formation and dynamics.
Rotational diffusion measurements of perylene sequestered within SDS and
CTAB micelles show that the viscosities within these confined environments is
higher than that of the corresponding alkane, and closer to that of the
corresponding length alkanol. This result is fully consistent with the literature and
chemical intuition, underscoring the balance of forces between ionic/polar

interactions within the palisades layer and van der Waals interactions within the

77

micelle interior. This balance of forces is sensitive to the composition of the
aqueous solution phase and the addition of optical acceptors to the micelle
solutions alters the balance of forces at the solution/micelle interface to the
extent of substantially altering the internal microviscosity of the micelles. Clearly
there is also a role played by the presence of n-butanol, a cosolvent used to
allow the introduction of perylene into the micellar solutions, and such effects
have been seen for similar n-alkanol additives previously. ‘8 Future
investigations will focus on the detailed reasons for the non-monotonic variation
in micelle microviscosity with increasing optical acceptor concentration. We
anticipate that the microscopic variations in micelle structure that result from the
balance between polar and dispersive forces in these systems will account for

our findings.

78

Literature Cited

(1) DelaCruz, J. L.; Blanchard, G. J. Joumal of Physical Chemistry B
2003, 107, 7102.

(2) Dutt, G. B. Journal of Physical Chemistry B 2002, 106.7398.

(3) Kelepouris, L.; Blanchard, G. J. Journal of Physical Chemistry B
2002, 106, 6600.

(4) Kelepouris, L.; Blanchard, G. J. Journal of Physical Chemistry B
2003, 107,1079.

(5) Klein, U. K. A.; Haar, H. P. Chemical Physics Letters 1978, 58, 531 .

(6) Maiti, N. C.; Krishna, M. M. G.; Britto, P. J.; Periasamy, N. Journal
of Physical Chemistry B 1997, 101, 1 1051 .

(7) Maiti, N. C.; Mazumdar, S.; Periasamy, N. Journal of Physical
Chemistry 1995, 99, 10708.

(8) Menger, F. M. Accounts of Chemical Research 1979, 12, 111.

(9) Quitevis, E. L.; Marcus, A. H.; Fayer, M. D. Journal of Physical
Chemistry 1993, 97, 5762.

(10) Rao, U. R. K.; Manohar, C.; Valaulikar, B. S.; lyer, R. M. Journal of
Physical Chemistry 1987, 91, 3286.

(11) Barzykin, A. V.; Tachiya, M. Journal of Chemical Physics 1995,
102, 3146.

(12) Choi, K. J.; Turkevich, L. A.; Loza, R. Journal of Physical Chemistry
1 988, 92, 2248.

(13) de Oliveira, H. P. M.; Gehlen, M. H. Chemical Physics 2003, 290,
85.

79

(14) De, 8.; Girigoswami, A.; Mandal, A. K. Spectrochimica Acta, Part A:
Molecular and Biomolecular Spectroscopy 2003, 59A, 2487.

(15) Ediger, M. D.; Domingue, R. P.; Fayer, M. D. Journal of Chemical
Physics 1984, 80, 1246.

(16) Kenney-Wallace, G. A.; Flint, J. H.; Wallace, S. C. Chemical
Physics Letters 1975, 32, 71 .

(17) Saha, D. 0.; Ray, K.; Misra, T. N. Spectrochimica Acta, Part A:
Molecular and Biomolecular Spectroscopy 2000, 56A, 797.

(18) Wistrom, C. A.; Jones, G. M.; Tobias, P. S.; Sklar, L. A. Biophysical
Journal 1996, 70, 988.

(19) Johansson, L. B. A.; Niemi, A. Journal of Physical Chemistry 1987,
91. 3020.

(20) Mattjus, P.; Molotkovsky, J. G.; Smaby, J. M.; Brown, R. E.
Analytical Biochemistry 1999, 268, 297.

(21 ) Stryer, L. Annual Review of Biochemistry 1978, 47, 819.

(22) Birch, D. J. S.; Suhling, K.; Holmes, A. S.; Salthammer, T.; lmhof,
R. E. Pure and Applied Chemistry 1993, 65, 1687.

(23) Ediger, M. D.; Fayer, M. D. Journal of Chemical Physics 1983, 78,
2518.

(24) Marcus, A. H.; Diachun, N. A.; Fayer, M. D. Journal of Physical
Chemistry 1992, 96, 8930.

(25) DeWitt, L.; Blanchard, G. J.; LeGoff, E.; Benz, M. E.; Liao, J. H.;
Kanatzidis, M. G. Journal of the American Chemical Society 1993, 115,
12158.

(26) Jha, B. K.; Patist, A.; Shah, D. O. Langmuir1999, 15, 3042.

80

(27) Dela Cruz, J. L.; Blanchard, G. J. Journal of Physical Chemistry A
2001, 105, 9328.

(28) Berr, S. S.; Caponetti, E.; Johnson, J. S., Jr.; Jones, R. R. M.;
Magid, L. J. Journal of Physical Chemistry 1 986, 90, 5766.

(29) Ben, S. 3.; Coleman, M. J.; Jones, R. R. M.; Johnson, J. 8., Jr.
Journal of Physical Chemistry 1986, 90, 6492.

(30) Ben, S. S. Journal of Physical Chemistry 1987, 91, 4760.
(31) Forster, T. Ann. Physik. 1948, 2,55.

(32) Jones, G. M.; Wofsy, C.; Aurell, C.; Sklar, L. A. Biophysical Journal
1999, 76,517.

(33) Keller, L.; Hussey, D. M.; Fayer, M. D. Journal of Physical
Chemistry 1996, 100, 10257.

(34) Benrraou, M.; Bales, B. L.; Zana, R. Journal of Physical Chemistry
B 2003, 107, 13432.

(35) Perche, T.; Auvray, X.; Petipas, C.; Anthore, R.; Rico, l.; Lattes, A.;
Bellissent, M. C. Journal de Physique H992, 2, 923.

(36) Dutt, G. B. Langmuir 2005, 21, 10391.

(37) Wu, 0.; Ma, R.; Zhou, B.; Shen, J.; Chan, K. K.; Woo, K. F.
Macromolecules 1996, 29, 228.

(38) Finger, K. U.; Marcus, A. H.; Fayer, M. D. Journal of Chemical
Physics 1994, 100,271.

(39) Gochanour, C. R.; Fayer, M. D. Journal of Physical Chemistry
1981 , 85, 1989.

81

(40) Chuang, T. J.; Eisenthal, K. B. Joumal of Chemical Physics 1972,
57, 5094.

(41) Klein, U. K. A.; Haar, H. P. Chemical Physics Letters 1979, 63, 40.
(42) Labhart, H.; Pantke, E. R. Chemical Physics Letters 1973, 23, 482.

(43) Hu, C.-M.; Zwanzig, R. Journal of Chemical Physics 1974, 60,
4354.

(44) Youngren, G. K.; Acrivos, A. Journal of Chemical Physics 1975, 63,
3846.

(45) Lipari, G.; Szabo, A. Biophysical Journal 1980, 30, 489.

(46) Debye, P. Polar Molecules. ln Chemical Catalog Co. New York,
1929.

(47) Edward, J. T. Journal of Chemical Education 1970, 47, 261.

(48) Goldie, S. N.; Blanchard, G. J. Journal of Physical Chemistry A
1999, 103, 999.

(49) Jiang, Y.; Blanchard, G. J. Journal of Physical Chemistry 1 994, 98,
6436.

(50) Jiang, Y.; Blanchard, G. J. Journal of Physical Chemistry 1995, 99,
7904.

(51) Palma, S.; Manzo, R. H.; Allemandi, D.; Fratoni, L.; Lo Nostro, P.
Colloids and Surfaces, A: Physicochemical and Engineering Aspects
2003, 212, 163.

(52) Zinsli, P. E. Chemical Physics 1977, 20, 299.

(53) Shinitzky, M.; Dianoux, A. G.; Gitler, G.; Weber, G. Biochemistry
1971, 10, 2106.

82

(54) Emert, J.; Behrens, C.; Goldenberg, M. Journal of the American
Chemical Society 1979, 101, 771.

(55) ’ Dutt, G. B. Journal of Physical Chemistry B 2004, 108, 3651.

83

 

Chapter 4

INVESTIGATING INTERNAL STRUCTURAL DIFFERENCES
BETWEEN MlCELLES AND UNILAMELLAR VESICLES OF
DECANOIC ACID/SODIUM DECANOATE

Introduction

Micelles and bilayer vesicle structures bear a loose structural
resemblance, but differ in fundamental ways in the details of their structure. .
Differences in the molecular organization of these structures are difficult to
compare directly because, in most cases, the molecules used to construct these
systems are different.

Micelles have been investigated extensively because they are capable of
encapsulating compounds in solution and because of their ability to mimic
biological systems. Properties such as micelle size, shape and permeability have
been found to depend sensitively on amphiphile identity and concentration. A
particularly useful tool for this work has been the interrogation of probe molecules
by fluorescence spectroscopy, where the dynamics of the probe molecule have
been related to system pressure, solvent viscosity, and the solubility of the
probe.“18

Unilamellar vesicle structures have found significance in immunological
studies, understanding cellular membrane structure and function, and drug

delivery. By far the most commonly used compounds for the formation of

unilamellar vesicles are phospholipids, a family of natural compounds that are

the principal constituents of plasma membranes. Vesicle structures can also be
formed by simpler amphiphilic compounds, such as octanoate, oleate and
decanoate.1M7 Decanoate micelles have been used for drug solubilization12 and
vesicles show potential for enzyme-catalyzed polymerizations.28 These
amphiphiles offer advantages over phospholipids, such as their dynamic

21 2229-31

behavior, ability to grow upon addition of more fatty acid,”2"”

autocatalytic formation,”-33

and structural simplicity, making them well suited to
fundamental investigations aimed at understanding organization in these
dynamic structures.

A further advantage that alkanoate amphiphiles offer is the ability to alter
the system morphology through adjustment of the solution pH. Sodium
decanoate solutions will form micelles at alkaline pH, provided the concentration
is above the CMC, and will form bilayers (vesicular) structures at intermediate

pH, provided the concentration is above the critical formation concentration,“

analogous to the phase transition reported in the more extensively studied

) 20.23.27 .34 d20.26.27 .35

caprylic acid (octanoate or oleic aci systems. The pH range
for vesicle formation has been reported as 6.8 - 7.8, with solutions of higher pH
existing in the micellar phase and solutions of lower pH resulting in phase
separation, with the formation of a precipitate.”23 If these spontaneously
assembled vesicles are extruded through a small-pore (100nm) filter, stable
vesicles of comparatively well defined size can be generated.“"3‘i'38

The fact that the same components form different structures allows for an

important comparison to be made directly. This is an issue of fundamental

85

 

significance, not only because of the passing structural similarity of micelles and
unilamellar vesicles, but because of the characteristic dimensions of these
structures. Micelles are expected to have a diameter on the order of 5 nm,
depending on the amphiphile used, while the unilamellar vesicles we use have
diameters on the order of 100 nm. It is important to consider the different radii of
curvature for the two types of structures and to understand from an experimental
standpoint the implications of the curvature of these systems on the internal
organization of the aliphatic regions. Indeed, the issue of curvature has been
discussed extensively in the context of phospholipid bilayer structures, but we
are not aware of any direct comparisons of internal organization and dynamics
between micelles and unilamellar vesicles comprised of the same amphiphiles.
We have sequestered perylene inside the sodium decanoate micelles and
unilamellar vesicles and have used fluorescence lifetime and anisotropy
measurements to obtain information on how the local environments differ
between these one and two layered systems. Perylene is a hydrophobic

chromophore that has been studied extensively,3740

making it an excellent
choice for sequestration in a micelle or vesicle structure. We have investigated
the decanoate micelle and vesicle systems to understand whether or not there is
a discernible difference in the local organization of the nonpolar regions of these
structures. Our data show that these structures do indeed produce different local

environments, with the unilamellar vesicle being of slightly lower effective

viscosity than the micelle.

86

Flgure 4.1 - The structures of perylene, sodium decanoate, and decanoic
acld

87

Experimental Section

Materials. Decanoic acid sodium salt (sodium decanoate, 98%) was
purchased from Fluka and used as received. Perylene was obtained from
Sigma-Aldrich, Inc. and used as received. These chemical structures can be
seen in Figure 4.1. Sodium phosphate, ethanol, and hydrochloric acid were
purchased from Fisher. The perylene concentration for all solutions was 10 11M.
For all decanoate solutions, perylene was first dissolved in an ethanol
solution,(10" M) then added to the decanoate solutions for solubility reasons.
Solution temperatures were maintained at 298 20.5 K (Neslab EX100-DD) for all
time-resolved fluorescence measurements. Micelle solutions were maintained at
a pH of ca. 9.3 (Orion Research digital ionanalyzer/501) using 150 mM
phosphate buffer, while vesicle solutions were lowered to a pH of ca. 6.8 by
adding 0.1 M HCI. All solutions were 100 mM in sodium decanoate,
approximately twice the critical micelle”41 and critical bilayerza'zs”28 forming
concentration.

Vesicle Formation. Vesicles were formed using the extrusion method.42
First, the solutions containing decanoate at the appropriate pH were subjected to
five freeze-thaw-vortex cycles. For each cycle the solution was immersed in
liquid nitrogen for five minutes, thawed in a hot water bath ca. 60°C for five
minutes, then vortexed for two minutes. The resulting solution was then passed
through polycarbonate filters with 100 nm diameter pores eleven times using a
Mini-Extruder system (Avanti Polar Lipids). The resulting unilamellar vesicles

were used within two days of extrusion.

88

Time-Correlated Single Photon Counting Measurements. Fluorescence
lifetime and anisotropy measurements of perylene in decanoate micelles and
decanoic acid/decanoate vesicles were made using a time-correlated single
photon counting (TCSPC) instrument that has been described in detail
previously.“3 The instrument description can be seen in Chapter 1. For the
experiments performed in this chapter, the excitation wavelength was set to 420
nm while the emission was collected at 470 nm.

Steady-State Spectroscopy. Absorption spectra were recorded using a
Cary model 300 double-beam UV-visible absorption spectrometer, with 1 nm
spectral resolution. Spontaneous emission spectra were acquired with a Spex
Fluorolog 3 spectrometer, with both excitation and emission monochromators set
to 3 nm resolution.

TEM Imaging. Unilamellar vesicle samples were stained with uranyl
acetate. A 5 pL drop was placed on formvar and carbon coated grids, allowed to
sit for several minutes, and the excess was removed with a filter paper. The
microscope was a JEOL 1OOCX (Japan) operated at an accelerating voltage of
100 W. A representative image of decanoate vesicles is shown in Figure 4.2a.

Cryo-TEM imaging. Cryogenic TEM images were acquired on our micelle
samples by Dow-Corning Analytical Solutions, Auburn Hills, MI. To prepare the
samples, ca. 2.3 mL of the micelle-containing solution was loaded on a holey
carbon film coated Cu TEM grid using a micropipette. The excess fluid on the
grid surface was removed by blotting the surface with a filter paper make an

aqueous thin film for the TEM. The grid was immersed in liquid ethane contained

89

 

Flgure 4.2 — (a) TEM Image of decanoic acid/decanoate unilamellar vesicles.
The average size Is ca. 100 nm diameter. (D) Cryo-TEM Image of decanoate
mlcelles Indicating a characteristic diameter of ca. 10 nm.

90

in a small vessel that itself was housed in a larger liquid nitrogen vessel. The
water film on the grid was vitrified to avoid water crystallization. The sample was
loaded into the TEM (JEOL JEM 2100F) and the morphology was observed
below 104 K. The digital images were acquired using a Gatan CCD camera
attached at the bottom of the TEM column and Digital Micrograph software. A

representative cryo-TEM image of the decanoate micelle is shown in Figure 4.2b.

91

Results and Discussion

The objective of this work is to better understand the differences between
the nonpolar regions of micelles and vesicles. Although micelles and vesicles
bear some structural resemblance, there are issues of curvature and the
characteristic length scale of any organization within these structures. To this
point, our ability to address these issues has been limited by the systems that are
capable of forming both types of structures. A recent report by Namani and i

Walde has shown the decanoic acid/sodium decanoate system to be capable of

 

forming either micelles or unilamellar vesicles, depending on solution properties
such as pH. They have reported that the pH range for vesicle formation in this
system is 6.4 to 7.8.28 The bilayer phase is stable when the pH of the system is
similar to the pK.a of the acid (decanoic acid pK.l ~ 4.83),“ due to hydrogen
bonding interactions between the protonated and deprotonated acid head
groups."’23“'2-"35 The difference between the pK.. of the acid and the actual pH at

19.213.28.3545-4‘3 and deals

which bilyer formation occurs has been discussed before,
with the variations between the pH of a polyanionic surface and the pH of the
bulk solution. Because both the micellar and vesicular systems are composed of
the same alkanoic acid, we can compare the dynamics of the perylene molecule
sequestered within each of these structural entities directly. The difference
between the lifetime and reorientation of the sequestered chromophore in the
micelle and vesicle structures will provide insight into the structural differences in

the aliphatic regions of these two structural forms. We consider the fluorescence

lifetime data first, to provide general information on the local environment(s) of

92

the perylene chromophores, then consider the fluorescence anisotropy data in
detail.

Fluorescence lifetime data. Fluorescence lifetime measurements of
perylene are useful in providing a qualitative picture of the local environment.
Perylene is less sensitive to either environmental polarity or the presence of
quenchers than some other polycyclic aromatic hydrocarbons (e. g. pyrene),“7 but
the information obtained from the data we report here on perylene still indicates
that the environments formed within the micelles and vesicles of decanoic
acid/decanoate are discemibly different. We recover a perylene fluorescence
lifetime of 5815 ns for the chromophore in decanoate/decanoic acid micelles, and
5942 ns for the chromophore in decanoate/decanoic acid unilamellar vesicles.
We note that for both systems we recover a single exponential decay population
decay, indicating rapid exchange between the distribution of environments that
are characteristic of each medium. The fact that the fluorescence lifetime is
shorter in the micelle than in the vesicle suggests that the environment in the
micelle is slightly more polar. This situation could result either from perturbations
to the micelle associated with amphiphile exchange with the solution or from the
location of perylene within the micelle. More likely is the difference in the
dielectric response of the interior of the micelle and vesicle structures. In the
micelle the influence of the surrounding solution dielectric is spatially uniform,
whereas it is anisotropic in the vesicle. There is a known relationship between
chromophore fluorescence lifetime and the presence of a dielectric gradient,‘“"51

and it is possible that this physical effect is responsible for our perylene lifetime

93

 

 

1.0 a

0.8

0.6

 

0.4

linear response (normalized)

0.2

 

Ohm

wavelength (nm)

 

linear response (normalized)

 

 

1 A l a l a l kl A I A I 24. I 1]

0.0'
360380400420440460480500520540560

wavelength (nm)

 

Figure 4.3 - (a) Normalized absorption and emission spectra of perylene In
100 mM decanoate/decanoic acid mlcelles (b) Normalized absorption and
emission spectra of perylene In 100 mM decanoate/decanoic acid vesicles.

94

 

data. This result may appear to be slightly at odds with the steady state
spectroscopic data (see Figure 4.3) showing essentially identical results for
micelles and vesicles. Given that the steady state spectra of perylene exhibit
such a weak dependence on solvent polarity, the steady state and lifetime results
are consistent with one another.

Fluorescence anisotropy measurements. In an effort to understand in
some level of detail the local environment of the probe molecule, we have
measured the fluorescence anisotropy dynamics of perylene. For the systems
we report on here, where the concentration of perylene within the micelles and
vesicles is low, with one perylene chromophore for every 100 to 200 micelles,
depending on the fraction of surfactant sequestered in the micelles.28 The
fluorescence anisotropy decay of the chromophore is associated with rotational
diffusion within the nonpolar region(s) of the micelles and vesicles. Rotational
diffusion is a well understood phenomenon and has been used extensively to
interrogate local organization in a variety of systems.‘°""‘“"‘"‘°"55 There is a well
established theoretical framework for the interpretation of reorientation data,‘°'5'*
5° and we consider several aspects of these data separately.

We obtain fluorescence depolarization data by exciting perylene with a
vertically polarized light pulse and collecting the emission decay transients
polarized parallel and perpendicular to the excitation polarization. These data

are combined to produce the induced orientational anisotropy function, a

description of which can be seen in Chapter 1.

95

W

 

 

 

 

 

 

 

 

1500
3'
E" 1000
E
sod
Delay Time (ps)
3 0.1
«1
V m
g l
‘9 l
515 000 I
.E '4 ‘
9 1 . , .
E) ilkm’Ill' I L‘ H“
(D 0.00 I I It‘ll-ind? 1"..J:_,_.I_I"1 a
0 309 100 gm
Delay Time (ps)

Figure 4.4 - (3) Experimental I.(t) and I 1(t) scans for perylene in decanoate
mlcelles, along with the instrumental response funcation. These data are
typical of those obtained for reorientation measurements. (b) Anisotropy
function, R(t), generated from the data shown In (a). The decay Is single
exponential and Is fit to the function R(t) = R(O) exp(-tlt-).

96

 

Because we are studying systems where the chromophore resides in a
confined environment, we need to consider the physical model that is best suited
to interpreting the experimental data. The choice of model will determine the
physical and chemical significance of the decay time constant(s) extracted from
the R(t) data. For chromophores tethered to a substantially larger entity, such as
a protein, or imbedded in a mono- or bilayer structure, the hindered rotor model
is appropriate. In this model the chromophore is attached to the larger assembly
at one point within the system, and its motion is considered in the context of the
conic volume it sweeps out. For the micelles and vesicles we consider here, the
hindered rotor model is not appropriate because the chromophore is not bound to
a specific point in or on the micelle or vesicle structure. Rather, perylene is able
to diffuse within the micelle or vesicle, and we thus treat this system in the
context of perylene reorienting in a quasi-bulk solvent. The model most
appropriate for this type of system is the modified Debye-Stokes-Einstein (DSE)
equation?“ which is covered in more detail in Chapter 1.

y37-40.60.61 ’ and

As noted above, perylene has been investigated extensivel
much about its reorientation dynamics are known. For perylene the
hydrodynamic volume is 225 A3,62 and S = 0.69. Because perylene is a
symmetric molecule, its transition dipole moments and rotational diffusion
Cartesian axes are coincident, and the Si «— So transition is known to be

polarized along its long in-plane axis, which is typically assigned as the x-axis.

For these experimental conditions, it has been established previously that a

97

"f

 

AT/I‘ (a.u.)

 

 

 

 

0 6000 12000
Delay Time(ps)
0.10 I
c: M,
0: 0.03 t'.
l “I
l l,
I 1W ,‘ L'I‘ H I l
o I'lllIiIIIIII‘IIHII‘I: I i -,.l .I‘ I
. I l 1 '1 1 I“
0'. 10'! 310'. 270'. I'O'O 0'0 1'0';

 

Delay Time (ps)

Figure 4.5 - (a) Experimental I.(t) and 11(1) scans for perylene In decanoate
vesicles, along wlth the Instrumental response funcation. These data are
typical of those obtained for reorientation measurements. (b) Anisotropy
function, R(t), generated from the data shown In (a). The decay Is single
exponential and Is fit to the function R(t) = R(O) exp(-t/r).

98

single exponential anisotropy decay corresponds to perylene reorienting as a
prolate rotor, with the time constant 10, = 1/GDZ.37'38'°° For the systems we have
examined here, we observe exclusively single exponential anisotropy decays,
indicating that perylene reorients as a prolate rotor in both micelles and vesicles
made of decanoic acid/decanoate. This is a significant finding when placed in
the context of previous work on perylene. This chromophore is known to exhibit
either a single or double exponential anisotropy decay, depending on
environment."“"63 The consistent finding of a single exponential decay in the
experiments we report here indicates that we can compare the data for perylene
in micelles and vesicles directly.

We consider next the appropriate treatment of the quantity f, which reflects
the frictional interactions between perylene and the constituents within the
nonpolar region of the micelles or vesicles. intuition may seem to suggest a
“slip” boundary condition for these systems because of their non-polar character,
but our earlier work on the reorientation of perylene in n-alkanes indicates that a
slip boundary condition is not appropriate.38 The experimental data for perylene
in n-alkanes, reorienting as a prolate rotor, are consistent with behavior
intermediate between the stick and slip limits. Experimentally, in the n-alkanes
perylene exhibits a viscosity dependence of 40 1 3 ps/cP for n-aikanes Cs — Ca,
and for C9 - C15, the viscosity dependence is measured as 12.7 :1: 1.5 ps/cP. We
believe that our data for micelles and vesicles can be treated in the context of a
12.7 ps/cP viscosity dependence owing to the length of the amphiphile aliphatic

chains. We measure experimentally to, = 268 :f: 3 ps for perylene in micelles,

99

corresponding to a viscosity of n = 21 (+3.1, -2.4) cP. For perylene reorienting in

decanoate vesicles, we measure r0, = 194 a: 10 ps, yielding n = 15 (+2.9, -2.3)
cP.

While there are few other data to compare with our results, the viscosity
values we infer are of some physical significance. First, we note that the
viscosity of the micelle is higher, outside of the experimental uncertainty, than
that of the vesicle. We anticipate that the extent of disorder and fraction of
gauche chain conformers in the micelle to be higher than that seen in the vesicle,
owing to the curvature of the micelle surface being greater than that of the
vesicle. TEM data (see Figure 4.2a) show that the decanoate unilamellar vesicles
are generally spherical in shape and are approximately 100 nm in diameter. The
larger diameter (micelles are ca. 10 nm diameter, Figure 4.2b) should result in a
decreased curvature. There exists a great deal of uncertainty in the viscosity of
micelle interiors as well as phospholipid bilayers. Previous studies of micelle
interiors proposed that the interiors were more rigid, with viscosities on the order
of 100 cP“ while others suggest that micelle interiors are quite fluid, with
viscosities on the order of 10 - 20 cP.""°“"66 Early work on phospholipid bilayer
structures likewise suggested a viscosity on the order of 100 cP,°7'°° but this
estimate has been shown to be relatively high and also quite dependent on the
constituents of the phospholipid bilayer.69 Koan and Blanchard have
recentiyexamined this issue and estimate the viscosity of a C14 phospholipid
bilayer to be in the range of 15 cP,‘39 a value similar to that we recover here for a

decanoate bilayer structure. We recognize that decanoate and a phospholipid

100

are very different compounds, both in terms of structure and polarity, but the
result we obtain for the decanoate bilayer structure is not inconsistent with these

findings.

 

101

Conclusions

We have studied the steady state and time resolved properties of the
chromophore perylene in both the micellar and vesicular structures of the
decanoate/decanoic acid/water system. The increase in the perylene
fluorescence lifetime in the decanoic acid/decanoate vesicle system is fully
consistent with the known behavior of the chromophore, and indicates that the
persistence time of any local organization is less than nanoseconds. The
fluorescence lifetime of perylene in the micelle is longer than in the vesicle, a
finding that may be related either to environmental polarity, disorder or to the
intrinsic differences in the dielectric environment of the micelle. We use
anisotropy decay data of the sequestered perylene to infer the viscosities of the
micelles (ca. 21 CF) and vesicles (ca. 15 GP). These findings are consistent with
other measurements of vesicle viscosity and suggest that the spatial confinement
imposed by the micelle curvature gives rise to less motional freedom within these
smaller structures than exists within the quasi-20 environment of the unilamellar

vesicles.

102

Literature Cited

(1) Jarzeba, W.; Walker, G. C.; Johnson, A. E.; Kahlow, M. A.; Barbara, P. F., J.
Phys. Chem, 1988, 92,7039.

(2) Visser, A. J. W. G.; Vos, K.; van Hoek, A.; Santema, J. S., J. Phys. Chem,
1988, 92, 759.

(3) Marcus, A. H.; Dichun, N. A.; Fayer, M. D., J. Phys. Chem, 1992, 96, 8930.

(4) Templeton, E. F. G.; Quitevis, E. L.; Kenney—Wallace, G. A., Journal of
Physical Chemistry, 1985, 89, 3238.

(5) Wirth, M. J.; Chou, S. H.; Piasecki, D. A., Analytical Chemistry, 1991, 63,
146.

(6) Streletzky, K.; Phillies, G. D. J., Langmuir, 1995, 11,42.
(7) Nandi, N.; Bagchi, B., J. Phys. Chem, 1996, 100, 13914.

(8) Maiti, N. C.; Krishna, M. M. G.; Britto, P. J.; Periasamy, N., J. Phys. Chem. B,
1997, 101, 11051.

(9) Matzinger, S.; Hussey, D. H.; Fayer, M. D., J. Phys. Chem. B, 1996, 102,
7216.

(10) Riter, R. E.; Willard, D. M.; Levinger, N. E., J. Phys. Chem. B, 1998, 102,
2705.

(11) Willard, D. M.; Riter, Ft. E.; Levinger, N. E., J. Am. Chem. 800., 1998, 120,
4151.

(12) Palma, S.; Manzo, Ft. H.; Allemandi, D.; Fratoni, L.; Lo Nostro, P., Colloids
and Surfaces, A: Physlcochemical and Engineering Aspects, 2003, 212, 163.

(13) Bhattacharyya, K.; Bagchi, B., J. Phys. Chem. A, 2000, 104, 10603.
(14) Cang, H.; Brace, D. D.; Fayer, M. D., J. Phys. Chem. 8,2001, 105, 10007.
(15) Hara, K.; Kuwabara, H.; Kajimoto, 0., J. Phys. Chem. A, 2001, 105, 7174.

(16) Chuang, T. J.; Eisenthal, K. B., Journal of Chemical Physics, 1972, 57,
5094.

(17) Mandal, D.; Sen, 8.; Bhattacharyya, K.; Tahara, T., Chem. Phys. Lett,
2002, 359, 77.

103

(18) Dutt, G. B.; Doraiswamy; Periasamy, N., Journal of Chemical Physics,
1991, 94, 5360.

(19) Gebicki, J. M.; Hicks, M., Nature (London, United Kingdom), 1973, 243,
232.

(20) Hargreaves, W. R.; Deamer, D. W., Biochemistry, 1978, 17,3759.

(21) Sacerdote, M. G.; Szostak, J. W., Proceedings of the National Academy of
Sciences of the United States of America, 2005, 102, 6004.

(22) Chen, l. A.; Szostak, J. W., Biophysical Journal, 2004, 87, 988.

(23) Morigaki, K.; Walde, P.; Misran, M.; Robinson, B. H., Colloids and Surfaces,
A: Physicochemical and Engineering Aspects, 2003, 213, 37.

(24) Chen, l. A.; Szostak, J. W., Proceedings of the National Academy of
Sciences of the United States of America, 2004, 101, 7965.

(25) Monnard, P.-A.; Deamer, D. W., Methods in Enzymology, 2003, 372, 133.

(26) Cistola, D. P.; Hamilton, J. A.; Jackson, 0.; Small, D. M., Biochemistry,
1988, 27, 1881.

(27) Walde, P.; Wick, R.; Fresta, M.; Mangone, A.; Luisi, P. L., Journal of the
American Chemical Society, 1994, 116, 11649.

(28) Namani, T.; Walde, P., Langmuir, 2005, 21, 6210.

(29) Kamp, F.; Zakim, D.; Zhang, F.; Noy, N.; Hamilton, J. A., Biochemistry,
1995, 34, 11928.

(30) Zhang, F.; Kamp, F.; Hamilton, J. A., Biochemistry, 1996, 35, 16055.
(31) Kleinfeld, A. M.; Chu, P.; Romero, C., Biochemistry, 1997, 36, 14146.

(32) Hanczyc, M. M.; Fujikawa, S. M.; Szostak, J. W., Science (Washington, DC,
United States), 2003, 302, 618.

(33) Oberholzer, T.; Wick, R.; Luisi, P. L.; Biebricher, C. K., Biochemical and
Biophysical Research Communications, 1995, 207, 250.

(34) Ekwall, P.; Mandell, L.; Fontell, K., Acta Chemica Scandinavica (1947-
1973), 1968, 22, 697.

104

(35) Haines, T. H., Proceedings of the National Academy of Sciences of the
United States of America, 1983, 80, 160.

(36) Hope, M. J.; Bally, M. B.; Webb, G.; Cullis, P. R., Biochimica etBiophysica
Acta, Biomembranes, 1 985, 812,55.

(37) Goldie, S. N.; Blanchard, G. J., Journal of Physical Chemistry A, 2001, 105,
6785.

(38) Jiang, Y.; Blanchard, G. J., Joumal of Physical Chemistry, 1994, 98, 6436.
(39) Jiang, Y.; Blanchard, G. J., Journal of Physical Chemistry, 1994, 98, 9411.
(40) Stevenson, S. A.; Blanchard, G. J., Langmuir, 2006, in review.

(41 ) McBain, J. W.; Salmon, 0. S., Journal of the American Chemical Society, '__
1920, 42,426. If

 

(42) MacDonald, R. C.; MacDonald, R. l.; Menco, B. P.; Takeshita, K.;
Subbarao, N. K.; Hu, L. R., Biochimica et biophysica acta, 1991, 1061, 297.

(43) DeWitt, L.; Blanchard, G. J.; LeGoff, E.; Benz, M. E.; Liao, J. H.; Kanatzidis,
M. G., Journal of the American Chemical Society, 1993, 115, 12158.

(44) Lide, D. R. CRC Handbook of Chemistry and Physics; CRC Press: Boca
Raton, FL, 1990-1991; Vol. 7fst Edition.

(45) Cistola, D. R; Atkinson, 0.; Hamilton, J. A.; Small, D. M., Biochemistry,
1986, 25, 2804.

(46) Apel, C. L.; Deamer, D. W.; Mautner, M. N., Biochimica et Biophysica Acta,
Biomembranes, 2002, 1559, 1.

(47) Karpovich, D. 8.; Blanchard, G. J., Journal of Physical Chemistry, 1995, 99,
3951.

(48) Drexhage, K. H., Journal of Luminescence, 1970, 1,2, 693.

(49) Girard, C.; Martin, O. J. F.; Dereux, A., Physical Review Letters, 1995, 75,
3098.

(50) Lukosz, W.; Kunz, R. E., Optics Communications, 1977,20, 195.

(51 ) Lukosz, W.; Kunz, R. E., Journal of the Optical Society of America, 1978,
67, 1607.

105

(52) Blanchard, G. J., Journal of Physical Chemistry, 1988, 92, 6303.

(53) Dela Cruz, J. L.; Blanchard, G. J., Journal of Physical Chemistry A, 2001,
105, 9328.

(54) Goldie, S. N.; Blanchard, G. J ., Book of Abstracts, 213th ACS National
Meeting, San Francisco, April 13-17, 1 997, ANYL.

(55) Kelepouris, L.; Blanchard, G. J., Journal of Physical Chemistry B, 2003,
107, 1079.

(56) Debye, P. Polar Molecules. In Chemical Catalog Co. New York, 1929.

(57) H0, C.-M.; Zwanzig, R., Journal of Chemical Physics, 1974, 60,4354.

(58) Perrin, F ., Journal de Physique et Ie Radium, 1936, 7, 1.

(59) Zwanzig, R.; Harrison, A. K., Journal of Chemical Physics, 1985, 83,5861.
(60) Edward, J. T., Journal of Chemical Education, 1970, 47, 261.

(61) Goldie, S. N.; Blanchard, G. J., Journal of Physical Chemistry A, 1999, 103,
999.

(62) Jiang, Y.; Blanchard, G. J., Journal of Physical Chemistry, 1995, 99, 7904.

(63) Piston, D. W.; Bilash, T.; Gratton, E., Journal of Physical Chemistry, 1989,
93, 3963.

(64) Povich, M. J.; Mann, J. A.; Kawamoto, A., Joumal of Colloid and Interface
Science, 1972, 41, 145.

(65) Shinitzky, M.; Dianoux, A. C.; Gitler, G.; Weber, G., Biochemistry, 1971, 10,
2106.

(66) Menger, F. M.; Jerkunica, J. M., Journal of the American Chemical Society,
1978, 100, 688.

(67) Cone, R. A., Nature: New biology, 1972, 236, 39.
(68) Poo, M.; Cone, R. A., Nature, 1974, 247, 438.

(69) Koan, M. M.; Blanchard, G. J., Journal of Physical Chemistry B, 2006, in
review.

106

 

Chapter 5

INVESTIGATING THE HYDROPHOBIC REGION OF MlCELLES
AND UNILAMELLAR VESICLES USING FLUORESCENCE
RESONANCE ENERGY TRANSFER

Introduction

Micelles have attracted considerable interest for a number of years due to
their many interesting properties, such as their size, shape, and permeability. In
addition, micelles have been shown to act as biomimetic and encapsulating
systems. Many studies of micelles have utilized time domain optical
spectroscopy to observe the dynamics of a probe molecule associated within the
micellar environment and to correlate the dynamics of the probe molecule with
that of the micelle as a function of viscosity, pressure, and probe solubility."18
Micelles and bilayer vesicle structures share many properties, such as the
amphiphilic nature of their constituents, but differ fundamentally in the details of
their structure. Unilamellar vesicles have also found use as biomimetic and
encapsulating systems and have been used in drug delivery as well as
immunological studies. Unlike micelles, which are typically composed of
surfactant molecules, unilamellar vesicles are usually formed fromphospholipids.
Since the molecules used to construct these structures are different, differences
in their molecular organization are difficult to compare directly.

Alkanoate amphiphiles, such as octanoate, oleate and decanoate, are

capable of forming micelles or vesicles depending on solution parameters, such

as pH.‘°’27 Sodium decanoate solutions will form micelles at alkaline pH,

107

provided the concentration is above the CMC, and will form bilayers (vesicular)
structures at intermediate pH, provided the concentration is above the critical

formation concentration,28 analogous to the phase transition reported in the more

d 2023.27.29 d20,26,27,30

extensively studied octanoic aci or oleic aci systems. The pH
range for vesicle formation has been reported as 6.8 - 7.8 for decanoic
acid/decanoate, with solutions of higher pH existing in the micellar phase and
solutions of lower pH resulting in precipitation.23'28 If the spontaneously-
assembled vesicles are extruded through a small-pore (100 nm) filter, stable
vesicles of comparatively well defined size can be generated.‘°'3"32 Since the
same molecular constituents can form either micelles or vesicles, we can make
relatively direct comparisons between the two structures.

For interrogating heterogeneous systems, it is possible to use fluorescent
compounds that will either partition selectively into a given phase or reside at the
interface between two phases. This environmental selectivity, when combined
with fluorescence resonance energy transfer (FRET) measurements, has been
used in the study of heterogeneous systems such as micelles, vesicles, and lipid
bilayers. 33‘“ The topology of these systems can significantly influence the
efficiency of energy transfer since the dipolar coupling process that mediates
energy transfer depends on both the average distance and relative orientation of
the molecules involved. Chromophore reorientation measurements can provide
information on the microenvironment(s) that exist within a heterogeneous

system. Micellar and vesicular systems are ideal candidates for energy transfer

experiments, and previous work in this area has examined energy transfer

108

between molecules in micelles and vesicles “"85"“ We are particularly
interested in the nonpolar region of micelles and vesicles, and understanding the
differences between these two structures that are comprised of the same
molecules.

We have sequestered perylene inside decanoate/decanoic acid micelles
and vesicles and have used the time-resolved fluorescence of this chromophore
to obtain information on how acceptor chromophores in the aqueous phase
interact with the donor. Perylene is a hydrophobic chromophore that has been
studied extensively, making it an appropriate choice for a probe to be
sequestered within the nonpolar region of micelles and vesicles. The ionic
acceptor chromophore, disodium fluorescein (DSF) has spectroscopic properties

well suited to its role as an optical acceptor.

109

Figure 5.1 — The structures of perylene, sodium decanoate, and decanoic
acid

110

Experimental Section

Materials. Decanoic acid sodium salt (sodium decanoate, 98%) was
purchased from Fluka and used as received. Perylene and disodium fluorescein
were obtained from Sigma-Aldrich, Inc and used as received. Structures for
these chemicals can be seen in Figure 5.1. Sodium phosphate, ethanol, and
hydrochloric acid were purchased from Fisher. For all solutions, perylene was
first dissolved in a 95% ethanol solution, (10“ M) then added to the decanoate
solutions for solubility reasons. The resulting solutions were ca. 10% ethanol,
90% aqueous micelle solution. For all time-resolved fluorescence
measurements, the solution temperatures were maintained at 298 :05 K
(Neslab EX100-DD). Micelle solutions were maintained at a pH of ca. 9.3 (Orion
Research digital ionanalyzer/501) using 150 mM phosphate buffer, while vesicle
solutions were lowered to a pH of ca. 7.0 by adding 0.1 M HCI. All solutions were

2357 and twice

100 mM sodium decanoate, approximately twice the critical micelle
the critical bilayerza'z‘a'28 formation concentration.

lime-Correlated Single Photon Counting Measurements. Fluorescence
lifetime and anisotropy measurements of perylene in decanoic acid/decanoate
micelle or vesicle containing solutions were made using a time-correlated single
photon counting (TCSPC) instrument that has been described in detail
previously,5a and a more complete description is available in Chapter 1. The

excitation wavelength was 420 nm and the sample emission was collected at 470

nm.

111

 

Vesicle Formation. Vesicles were formed using the extrusion method.42
First, the solutions containing decanoate (100 mM) at pH 7.0 were subjected to
five freeze-thaw-vortex cycles, with each cycle consisting of freezing the sample
in liquid nitrogen for five minutes, then heating the sample in a hot water bath
(ca. 60° C) for five minutes, followed by vortexing the sample for two minutes.
The resulting solution was then passed through polycarbonate filters with 100 nm
diameter pores eleven times using a Mini-Extruder system (Avanti Polar Lipids).
The resulting unilamellar vesicles were used within two days of extrusion.

TEM Imaging. Unilamellar vesicle samples were stained with uranyl
acetate. A 5 [1L drop was placed on formvar and carbon coated grids, allowed to
sit for several minutes, and the excess was removed with a filter paper. The
microscope was a JEOL 1000X (Japan) operated at an accelerating voltage of
100 W. A representative image of decanoate vesicles is shown in Figure 5.2a.

Cryo-TEM imaging. Cryogenic TEM images were acquired on our micelle
samples by Dow-Corning Analytical Solutions, Auburn Hills, Mi. To prepare the
samples, ca. 2.3 mL of the micelle-containing solution was loaded on a holey
carbon film coated Cu TEM grid using a micropipette. The excess fluid on the
grid surface was removed by blotting the surface with a filter paper make an
aqueous thin film for the TEM. The grid was immersed in liquid ethane contained
in a small vessel that itself was housed in a larger liquid nitrogen vessel. The
water film on the grid was vitrified to avoid water crystallization. The sample was
loaded into the TEM (JEOL JEM 2100F) and the morphology was observed

below 104 K. The digital images were acquired using a Gatan CCD camera

112

‘r
‘ ,er.
1205' .

 

Figure 5.2 - (a) TEM Image of decanoic acid/decanoate unilamellar vesicles.
The average size is ca. 100 nm diameter. (b) Cryo-TEM Image of decanoate
mlcelles (dark features) indicating a characteristic diameter of ca. 10 nm.

113

attached at the bottom of the TEM column and Digital Micrograph software. A

representative cryo-TEM image of the decanoate micelle is shown in Figure 5.2b.

114

 

Results and Discussion

The objective of this work is to better understand the differences between
the hydrophobic regions of micelles and vesicles. Micelles and vesicles bear
some structural similarity, but there are differences in the curvature and
characteristic length scale of any organization within these structures. To this
point, our ability to address these differences has been limited by the number of
systems that are capable of forming both micelle and vesicle structures. A recent
report by Namani and Walde has shown the decanoic acid/sodium decanoate
system to be capable of forming either micelles or vesicles, depending on the
solution pH.23 They have reported that the pH range for vesicle formation is
between 6.4 and 7.8. The bilayer-forming phase is stable when the pH of the

)f9due

system is similar to the pK,, of the acid (the pK,, for decanoic acid is 4.83
to hydrogen bonding interactions between the protonated and deprotonated acid
head groups.”'2"27'3° The difference between the pK. of the acid and the actual

192828305051 With a

pH at which bilyer formation occurs has been studied before,
significant focus on the deviation between the pH of the bulk solution and the pH
of a polyanionic surface. Because both the micelle and vesicle systems are
composed of the same alkanoic acid, we can compare the dynamics of the
perylene molecule sequestered within each of these structural entities directly.
Differences in fluorescence lifetime and reorientation time for perylene
sequestered in micelles and vesicles provides insight into the structural

differences in the aliphatic regions of these two structural forms. We first

consider the fluorescence lifetime data to provide a qualitative comparison of the

115

local environment(s) characteristic of these two structures, then consider the
fluorescence anisotropy data for each to gain further insight.

Fluorescence lifetime data. Fluorescence lifetime measurements of
perylene are useful in providing a qualitative picture of the local environment.
The concentration of optical donor in the micelles is such that we estimate there
to be a single perylene chromophore in approximately 0.5% of the micelles, while
for vesicles, the chromophorezvesicle ratio is close to unity. The optical acceptor
concentrations we use are such that in a homogeneous solution, the average
distance between donor and acceptor would be 150 A or more. Under these
experimental conditions, very little donor-acceptor excitation transfer will be seen,
and we will observe acceptor concentration-dependent changes in the donor
fluorescence lifetime only if the optical acceptor interacts preferentially with the
micelles or vesicles.

The transfer of energy between molecules can occur by a number of
mechanisms, ranging from dipolar coupling to collisional quenching. We are
concerned in this work with the former phenomenon because of the characteristic
length scale over which it operates. The transfer of energy by dipolar coupling
has been described by Fbrster, 62
km: “2RD J88x10‘7IA0/2f0(‘7)d17=£2—I£ 59 5.1

T36 n4 R
where km is the rate constant for energy transfer between donor and acceptor.

The K term is a function of the relative orientation of the donor and acceptor

transition dipole moments and, for our experimental conditions «2>= 2/3. The

116

term k,D is the donor radiative decay rate constant in the absence of optical

acceptor, n is the refractive index and R is the intermolecular donor-acceptor
distance. The integral term is a measure of the spectral overlap between the
emission spectrum of the optical donor and the absorption spectrum of the

acceptor. The spectral overlap term, combined with the several constants on the
right hand side of Eq. 5.1 is typically referred to as the critical radius, R0, a
quantity which serves as an effective gauge of the operating distance of this

dipolar coupling process. While the term R0 will vary for each donor-acceptor

pair, for most organic chromophore pairs, R0 ~ 50 A. “555253 Based on the
spectral profiles of perylene emission and DSF absorption, and the extinction and
fluorescence quantum yield data on these molecules, we estimate R0 ~ 54 A for
the perylene/DSF pair. We excite perylene at 420 nm and measure its
fluorescence lifetime at ca. 470 nm, without interference from acceptor emission.
The measurement of donor lifetime as a function of acceptor concentration is a
direct means of evaluating variations in km, since this rate constant will operate
in an additive manner with the intramolecular decay rate constant(s) that govern
the depopulation of the donor excited state. Since the concentration of perylene
is held constant at 10'5 M, energy transfer between two donor molecules should
be negligible because of the small fraction of micelles that contain a perylene
molecule. Even if this process were significant, it would appear as a

depolarization of the perylene fluorescence, but not a diminution of its lifetime. 6‘

117

 

 

 

 

 

 

2000
1500
£1000
l:
"5
500 i
.J‘
Woo
Delay Time (ps)
600
3
E;
I: 300
'2
0 than... .
0 6000 12000
Delay Time (ps)

Figure 5.3 - (a) Experimean scans for perylene in decanoate vesicles,
along with the Instrumental response function. (b) Experimental scans for
perylene in decanoate vesicles and 10‘3 M DSF present. These data are
typical of those obtained for lifetime measurements.

118

2400

 

 

 

 

A 1 800
3.

3
t 1200
'2

600

0
O 5000 1 0000 1 5000
Delay Time (ps)

Figure 5.4 - Experimental scans for perylene In decanoate mlcelles, along
with the instrumental response function.

119

 

 

 

 

 

 

Acceptor T111 (PS) T111 (133)
Concentration (M)
0 5942 :l: 10 -
10'5 5977 :I: 19 -
10“ 5806 9. 9 -
10'3 4723 :I: 12 659 :l: 16

 

 

 

 

 

 

 

 

b
Acceptor rm (ps)
Concentration (M)
o 5815 :1: 1o
10'5 5681 :I: 18
10" 5530 :I: 19
10°3 4196 :I: 12

 

 

 

 

 

Table 5.1 - (a) Fluorescence lifetime data for perylene sequestered in
decanoate vesicles. (b) Fluorescence lifetime data for perylene sequestered
in decanoate/decanoic acld mlcelles. The data reported here are the
averages of 6 Independent determinations, and the uncertainties reported
are :1: 10.

120

We first consider the interactions between perylene and DSF in decanoic
acid/decanoate unilamellar vesicles. As shown in Table 5.1a, the fluorescence
lifetime of perylene decreases as the DSF concentration increases. In addition to
the decrease in fluorescence lifetime, the increasing acceptor concentration also
affects the functionality of the perylene decay. For 10'3 M DSF, we observe a
two-component perylene fluorescence lifetime. Representative experimental
scans for the vesicle system can be seen in Figure 5.3, while micelle scans are
presented in Figure 5.4. The data for both systems is shown in Table 5.1. We
understand this behavior in the context of interactions between the acceptor
chromophore and the vesicle, and will return to this discussion after considering
the micelle data.

We compare the vesicle data to those that we have obtained for perylene
and DSF in decanoate micelles. As the concentration of acceptor increases, we
observe a decrease in the fluorescence lifetime of perylene. For 10'3 M DSF
concentration, however, we do not observe a change in the functionality of the
perylene population decay. The fact that we observe a change functionality of
the decay for perylene in vesicles but not in micelles is interesting and points to
the difference between the surface charge of the two structures. Also of interest
are the exponential prefactors for the perylene double exponential decay in
vesicles. For 10'3 M DSF in the vesicle-containing solution, the fast lifetime
component (1: = 659 ps) represents 38% of the total, with the longer decay (r:

4723 ps) accounting for 62% of the total decay intensity. The weighted average

121

of these lifetimes, the value that we get is somewhat faster than the perylene
lifetime found for the micelle system with 10'3 M acceptor present. The finding of
the two-component donor decay for vesicle systems at high acceptor
concentration and the one component donor decay in micelle systems could be
the result of the different relative abundances of the micelles and vesicles.

In addition to qualitative insight into excitation transport between the
interior and exterior region of micelles and vesicles, the lifetime data can also
provide somewhat more quantitative insight into the properties of the interfacial
region. We consider that acceptors form a complex with the micelle. In this
picture, the acceptors are either in very close spatial proximity to the donor when
interacting with the micelle, or too far distant to transfer excitation efficiently when
not interacting with the micelle. This model is clearly an oversimplification, but it
allows for the ready use of the Forster equation in the estimation of association
constants for these systems.

It is first important to consider the approximate stoichiometry of the donor,
acceptor and micelles in our solutions. As noted above, the perylene donor
concentration is such that there is one donor per ~200 micelles. As we increase
the acceptor concentration from 10'5 M to 10'3 M, the acceptor-to-micelle
stoichiometry changes from one acceptor per ~ 200 micelles to one acceptor for
~2 micelles. Because of the perylene:micelle ratio of 1:200, for a 10'5 M acceptor
concentration, there is one acceptor per ~200 micelles, so only 0.0025% of the
micelles will have both a donor sequestered within and an acceptor complexed to

them. Under these conditions, there will be essentially no donor-containing

122

micelles that are complexed to more than one acceptor. We can thus
approximate safely that for low acceptor concentrations, the equilibrium process
we monitor with our donor lifetime measurements corresponds to interactions
between single donors and single acceptors.

Before considering the information content of these data, we need to
evaluate the sizes of the micelles under investigation. The cryo-TEM data on our
micelles (Figure 5.2b) indicate that the micelles are in the sub-10 nm range, but
we are not able to extract precise average diameters. It is important to consider
that these micelles have experienced vitrification, so it is not unreasonable to
expect that there has been some structural perturbation prior to acquisition of the
images. The larger point of these images is that we do indeed have nanoscale
micelles present in our solutions, and with their existence established, we will use
the more precise estimates of micelle size that have been reported elsewhere.
Specifically, the radius of a sodium decanoate micelle is taken to be 23 A.65

The lifetimes we measure are treated as weighted averages of the
fluorescence lifetime of perylene in micelles complexed with one acceptor and
the lifetime of perylene in an uncomplexed micelle. We take the experimentally
determined fluorescence lifetime as the weighted average of the free form are, =
(k,")'1 and the complexed form, where top... = (k,D + km)", with koA being
calculated for R =23 A and R; as given above.

+n

= n cplecplx

To freeTfree

bs
5.2
,, [MA]

=_["_”]__ n =___
free [M]+[MA]’ CP/X [M]+[MA]

123

From these calculations, we recover the fractional contributions from the
free and complexed forms of the perylene-containing micelles. For the acceptor

association with the micelles, the relevant equilibrium process is:

= [MA] = neplx
[M HA] n “[14]

 

M+A K MA K 5.3

 

We know [M] and [A] from solution composition, and determine the ratio of

[MAJ/[M] experimentally from the lifetime measurements. We determine the
quantities m,“ and new, (Eq. 5.2), and use them to calculate K. We find that for

DSF present at 10‘5 M in decanoate vesicles, K is not measurable. If we
increase the DSF concentration to 10“ M, we recover K ~ 230 M". For DSF
present at 10'5 M in micelles, we find that K ~ 2300 M". If we increase the DSF
concentration to 104 M and 10"3 M, we recover ~ 519 M'1 and ~ 389 M'1
respectively. It is possible that the formation constant for the 10" M and 10—3 M
system is not markedly different from that of the 10‘5 M solution, but rather the
difference between the values it due to more than one acceptor molecule

complexing to the micelle. For a multiple equilibrium process,

M+ A$MA+ A:'_(_2—\_MA2 K: K,K2 5.4
From the micelle experiment with 10'5 M acceptor, K1 ~ 2000, and for the 10‘4 M
acceptor solution, K1K2 ~ 500 with the 10'3 M DSF solution, K1K2 ~400. This
leads to K2 values of ~ 0.25 and ~ 0.2, respectively. While it is normal for the
equilibrium constants to diminish monotonically with the addition of complexing

species, the difference between K1 and K2 for acceptor/micelle complexation is

substantial. It is possible that the facile translational motion of the acceptor on

124

 

 

the micelle or vesicle “surface” as well as the size of the acceptor molecules give
rise to a significant contribution from steric effects for the association of multiple
acceptors with a given micelle or vesicle.

Rotational diffusion within the micelles. We have studied the rotational
diffusion dynamics of perylene within the micelles and vesicles as a function of
acceptor concentration in order to understand the influence of the acceptor
molecules on the motional freedom of the sequestered donor. Rotational
diffusion has been used extensively to interrogate local organization in solutions,
because of the well developed theoretical framework for interpretation of the data
and the relatively high information content of the experimental signals. “MM
Our data indicate that the acceptor molecules influence donor motion differently
in micelles and vesicles.

We obtain fluorescence depolarization data by exciting perylene with a
vertically polarized light pulse and collecting the emission decay transients
polarized parallel and perpendicular to the excitation polarization. These data
are combined to produce the induced orientational anisotropy function, R(t),
which is discussed in Chapter 1, equation 1.2. The chemical information
contained in the data lies in the decay time constant(s) of R(t), and its zero-time
value. We note that donor-donor excitation transport can give rise to
fluorescence depolarization, the same physical phenomenon that we use to
characterize the molecular motion of perylene sequestered within micelles. The
D-A excitation transport data (vide supra) point to modest donor-acceptor

excitation transport, and given the low perylene concentrations used (~1 perylene

125

 

per 200 micelles), we believe that fluorescence depolarization resulting from
donor-donor transport does not contribute measurably to our data. Experimental
scans for perylene in decanoate vesicles as well as decanoate micelles can be
seen in Figures 5.5 and 5.6, respectively.

Because we are studying heterogeneous systems, it is fair to consider
which model is most appropriate for the treatment of the anisotropy data. For
chromophores tethered to a substantially larger entity, such as a protein, or
imbedded in a mono- or bilayer structure, the hindered rotor model is
appropriate.”69 In this model the chromophore is bound at one point within the
system and its motion is considered in the context of the conic volume it sweeps
out. For the micellar systems we are concerned with here, the hindered rotor
model is not appropriate because the chromophore is not bound to a specific
point in or on the micelle. Rather, perylene is free to diffuse within the micelle,
and we treat this system in the context of perylene reorienting in a quasi-bulk
solvent. The model most appropriate for this type of system is the modified
Debye-Stokes-Einstein (DSE) equation, 7° which is explained in Chapter 1.

For perylene the hydrodynamic volume is 225 A3, T = 300 K, S = 0.69 (prolate
rotor based on single exponential decay). "'72 Our experimental anisotropy data
are all single exponential decays, consistent with perylene reorienting as a
prolate rotor within the micelles. Perylene has been shown previously to reorient
either as a prolate rotor (single exponential anisotropy decay) or as an oblate

rotor (double

126

 

AT/l' (a.u)

2000

 

1000 P

o. ' A

0 5000 10000 15000 20000
Delay Time(ps)

 

 

0.2

R(t)

.11 '
0.1 '

1 "1.1

0.0 I r.l.l!ll£!i!lli1 A ll . 11111. Illa:

 

 

0 200 400 600 800 100012001400
DelayTime(ps)

L

Figure 5.5 - (a) Experimental I.(t) and I i(t) scans for perylene In decanoate
vesicles, along with the Instrumental response funcation. These data are
typical of those obtained for reorientation measurements. (b) Anisotropy
function, R(t), generated from the data shown in (a). The decay is single
exponential and is fit to the function R(t) = R(O) exp(-tlr).

127

2500
2000

““1

1000

MN (a.u.)

 

q __l L
Delay Time (ps)

 

 

Flit)
O
___

 

0.00 "1.1.1th

 

 

Delay Time (ps)

Figure 5.6 - (a) Experimental I.(t) and Mt) scans for perylene in decanoate
mlcelles, along with the instrumental response funcation. These data are
typical of those obtained for reorientation measurements. (b) Anisotropy
function, R(t), generated from the data shown in (a). The decay is single
exponential and is fit to the function R(t) = R(0) exp(-tlr).

128

conditions, experimental data for perylene in n—alkanes are consistent with
behavior intermediate between the stick and slip limits. From the modified DSE
model, in the stick limit we calculate the quantity :an = 78 ps/cP, and in the slip
limit we calculate 1an = 5.8 ps/cP. Experimentally we know from earlier work on
perylene in n-alkanes longer than Ca, 1an = 12.7 :1: 1.5 ps/cP, intermediate
between the stick and slip limit, and we use this experimental value to estimate
the viscosity of the micellar media.‘3'7"7‘ For perylene in the sodium
decanoate/decanoic acid micelles in the absence of acceptor molecules, the
effective viscosity is 21 cP and with the addition of anionic DSF acceptor, the
effective viscosity of the interior of the micelle decreases to 10 cP for 10'3 M DSF
(see Table 5.2a). The decrease in apparent viscosity with the addition of
acceptor indicates that the solution phase ionic species interact with the micelle
in such a way at to make the interior of the micelle more fluid. It is interesting that
the largest decrease in the effective viscosity of the micelle interior occurs upon
the initial addition of acceptor molecules. This indicates that the presence of DSF
in the decanoate/decanoic acid micelles is somehow perturbing the micellar
system, an effect which might result from the properties of the acceptor molecule.
For sodium decanoate/decanoic acid vesicles, we do not observe
analogous monotonic trends with increasing acceptor concentration (see Table
5.2b). For the native vesicle, in the absence of any optical acceptor, we recover

a viscosity of 15 cP for the hydrophobic region. As anionic DSF acceptor is

129

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Acceptor Cancentration Ton (ps) '1 (CF)
0 1 94 :1: 10 15
105 141.7 :1: 3 1 1
104 142.4 :I: 11 11
1 0-3 1 86.8 :1: 3 14
Acceptor (fancentration Ton fl (cP)
0 268 :1: 6 21
10-5 175 1 4 14
10-4 144.3 :1: 7 1 1
1 08 132.2 :1: 8 10

 

 

 

 

 

Table 5.2 - (a) Rotational diffusion time constants of perylene sequestered
In the nonpolar region of decanoic acid/decanoate vesicles (b) Rotational
diffusion time constants of perylene sequestered inside decanoate
mlcelles. The data reported here are the averages of 6 Independent
determinations, and the uncertainties reported are :1: lo.

130

added to the vesicle-containing solution, the viscosity of the nonpolar region
ranges from 11.0 CF to 14.0 CF, but there is no clear trend in these data.

From these effective viscosity data, we can infer the mobility of perylene
sequestered within the vesicle walls. The model used to relate the translational
diffusion coefficient to solution parameters such as viscosity is the Stokes-

Einstein equation"
D=—— (4)

where D is the translational diffusion coefficient of perylene and r is the radius of
a sphere of volume 225 A3. For the vesicle solution without any DSF present, the
viscosity was 15 cP, we recover a translational diffusion coefficient of 4.2 x10'7
cmzs“. This value is an order of magnitude larger than the diffusion coefficients
typically observed for lipid systems.5 The difference between our value and
those observed for lipid systems is most likely due to the size of our vesicles, as
well as radius of curvature of our systems. We also need to consider the validity
of the Stokes-Einstein equation. The Stokes-Einstein equation is valid under
ideal conditions for homogenous solutions. It also assumes that the diffusing
molecule is a sphere. The solutions that we are examining are obviously not

homogenous, and perylene is not a perfect sphere.

131

Conclusions

We have examined the fluorescence lifetime and reorientation dynamics
of perylene sequestered in micelle and vesicle structures formed from sodium
decanoate/decanoic acid solutions in the presence of varying concentrations of
the ionic optical acceptor DSF. The decrease in perylene lifetime in the presence
of DSF is consistent with dipolar excitation transport. We use anisotropy decay
data to compare the viscosities of the hydrophobic regions of micelles and
vesicles in the presence of acceptor molecules and find that the rigidity of the
micellar structures decreases with the increase in ionic strength, while no change

is noted in the vesicle structures.

132

 

Literature Cited

(1) Jarzeba, W.; Walker, G. C.; Johnson, A. E.; Kahlow, M. A.;
Barbara, P. F. J. Phys. Chem. 1988, 92, 7039.

(2) Visser, A. J. W. G.; Vos, K.; van Hoek, A.; Santema, J. S. J. Phys.
Chem. 1988, 92, 759.

(3) Marcus, A. H.; Dichun, N. A.; Fayer, M. D. J. Phys. Chem. 1992,
96. 8930.

(4) Templeton, E. F. G.; Quitevis, E. L.; Kenney-Wallace, G. A. Journal
of Physical Chemistry 1985, 89, 3238.

(5) Vaz, W. L. C.; Criado, M.; Madeira, V. M. C.; Schoellmann, G.;
Jovin, T. M. Biochemistry 1982, 21 , 5608.

(6) Streletzky, K.; Phillies, G. D. J. Langmuir1995, 11,42.

(7) Nandi, N.; Bagchi, B. J. Phys. Chem. 1996, 100, 13914.

(8) Maiti, N. C.; Krishna, M. M. G.; Britto, P. J.; Periasamy, N. J. Phys.
Chem. B1997, 101, 11051.

(9) Matzinger, S.; Hussey, D. H.; Fayer, M. D. J. Phys. Chem. B 1998,
102, 7216.

(10) Riter, R. E.; Willard, D. M.; Levinger, N. E. J. Phys. Chem. B 1998,
102, 2705.

(11) Willard, D. M.; Riter, R. E.; Levinger, N. E. J. Am. Chem. Soc.
1998, 120, 4151 .

(12) Palma, S.; Manzo, R. H.; Allemandi, D.; Fratoni, L.; Lo Nostro, P.
Colloids and Surfaces, A: Physicochemical and Engineering Aspects
2003, 212, 163.

133

(13) Pal, S. K.; Mandal, D.; Sukul, D.; Bhattacharyya, K. Chemical
Physics 1999, 249, 63.

(14) Cang, H.; Brace, D. D.; Fayet, M. D. Journal of Physical Chemistry
B2001, 105, 10007.

(15) Hara, K.; Kuwabara, H.; Kajimoto, O. J. Phys. Chem. A2001, 105,
7174.

(16) Chuang, T. J.; Eisenthal, K. B. Journal of Chemical Physics 1972,
57, 5094.

(17) Mandal, D.; Sen, 8.; Bhattacharyya, K.; Tahara, T. Chem. Phys.
Lett. 2002, 359, 77.

(18) Dutt, G. B.; Doraiswamy; Periasamy, N. Journal of Chemical
Physics 1991, 94, 5360.

(19) Gebicki, J. M.; Hicks, M. Nature (London, United Kingdom) 1973,
243, 232.

(20) Hargreaves, W. R.; Deamer, D. W. Biochemistry1978, 17,3759.

(21) Sacerdote, M. G.; Szostak, J. W. Proceedings of the National
Academy of Sciences of the United States of America 2005, 102, 6004.

(22) Chen, l. A.; Szostak, J. W. Biophysical Joumal 2004, 87, 988.

(23) Morigaki, K.; Walde, P.; Misran, M.; Robinson, B. H. Colloids and
Surfaces, A: Physicochemical and Engineering Aspects 2003, 213, 37.

(24) Chen, l. A.; Szostak, J. W. Proceedings of the National Academy of
Sciences of the United States of America 2004, 101, 7965.

(25) Monnard, P.-A.; Deamer, D. W. Methods in Enzymology 2003, 372,
133.

134

 

(26) Cistola, D. P.; Hamilton, J. A.; Jackson, 0.; Small, D. M.
Biochemistry 1988, 27, 1881 .

(27) Walde, P.; Wick, R.; Fresta, M.; Mangone, A.; Luisi, P. L. Journal of
the American Chemical Society 1994, 116, 11649.

(28) Namani, T.; Walde, P. Langmuir 2005, 21 , 6210.

(29) Ekwall, P.; Mandell, L.; Fontell, K. Acta Chemica Scandinavica
(1947-1973) 1968, 22, 697.

(30) Haines, T. H. Proceedings of the National Academy of Sciences of
the United States of America 1983, 80, 160.

(31) Hanczyc, M. M.; Fujikawa, S. M.; Szostak, J. W. Science
(Washington, DC, United States) 2003, 302,618.

(32) Hope, M. J.; Bally, M. B.; Webb, G.; Cullis, P. R. Biochimica et
Biophysica Acta, Biomembranes 1985, 812, 55.

(33) DelaCruz, J. L.; Blanchard, G. J. Journal of Physical Chemistry B
2003. 107, 7102.

(34) Dutt, G. B. Journal of Physical Chemistry B 2002, 106, 7398.

(35) Kelepouris, L.; Blanchard, G. J. Journal of Physical Chemistry B
2002, 106, 6600.

(36) Kelepouris, L.; Blanchard, G. J. Journal of Physical Chemistry B
2003, 107, 1079.

(37) Klein, U. K. A.; Haar, H. P. Chemical Physics Letters 1978, 58,531.

(38) Maiti, N. C.; Krishna, M. M. G.; Britto, P. J.; Periasamy, N. Journal
of Physical Chemistry B 1997, 101, 11051.

135

 

(39) Maiti, N. C.; Mazumdar, S.; Periasamy, N. Journal of Physical
Chemistry 1 995, 99, 10708.

(40) Menger, F. M. Accounts of Chemical Research 1979, 12, 111.

(41) Quitevis, E. L.; Marcus, A. H.; Fayet, M. D. Journal of Physical
Chemistry 1993, 97, 5762.

(42) MacDonald, R. C.; MacDonald, R. |.; Menco, B. P.; Takeshita, K.;
Subbarao, N. K.; Hu, L. R. Biochimica et biophysica acta 1991, 1061, 297.

(43) Barzykin, A. V.; Tachiya, M. Journal of Chemical Physics 1995,
102, 3146.

(44) Choi, K. J.; Turkevich, L. A.; Loza, R. Journal of Physical Chemistry
1988. 92,2248.

(45) de Oliveira, H. P. M.; Gehlen, M. H. Chemical Physics 2003, 290,
85.

(46) De, S.; Girigoswami, A.; Mandal, A. K. Spectrochimica Acta, Part A:
Molecular and Biomolecular Spectroscopy 2003, 59A, 2487.

(47) Ediger, M. D.; Domingue, R. P.; Fayer, M. D. Journal of Chemical
Physics 1984, 80, 1246.

(48) Kenney-Wallace, G. A.; Flint, J. H.; Wallace, S. C. Chemical
Physics Letters 1975, 32, 71 .

(49) Saha, D. C.; Ray, K.; Misra, T. N. Spectrochimica Acta, Part A:
Molecular and Biomolecular Spectroscopy 2000, 56A, 797.

(50) Wistrom, C. A.; Jones, G. M.; Tobias, P. S.; Sklar, L. A. Biophysical
Journal 1996, 70, 988.

(51 ) Johansson, L. B. A.; Niemi, A. Journal of Physical Chemistry 1987,
91 , 3020.

136

 

(52) Mattjus, P.; Molotkovsky, J. G.; Smaby, J. M.; Brown, R. E.
Analytical Biochemistry 1 999, 268, 297.

(53) Stryer, L. Annual Review of Biochemistry 1978, 47, 819.

(54) Birch, D. J. S.; Suhling, K.; Holmes, A. S.; Salthammer, T.; lmhof,
R. E. Pure and Applied Chemistry 1993, 65, 1687.

(55) Ediger, M. D.; Fayer, M. D. Journal of Chemical Physics 1983, 78,
2518.

(56) Marcus, A. H.; Diachun, N. A.; Fayer, M. D. Journal of Physical
Chemistry 1992, 96, 8930.

(57) McBain, J. W.; Salmon, C. S. Journal of the American Chemical
Society 1 920, 42, 426.

(58) DeWitt, L.; Blanchard, G. J.; LeGoff, E.; Benz, M. E.; Liao, J. H.;
Kanatzidis, M. G. Journal of the American Chemical Society 1993, 115,
12158.

(59) Lide, D. R. CRC Handbook of Chemistry and Physics; CRC Press:
Boca Raton, FL, 1990-1991; Vol. 71st Edition.

(60) Cistola, D. R; Atkinson, D.; Hamilton, J. A.; Small, D. M.
Biochemistry 1986, 25, 2804.

(61) Apel, C. L.; Deamer, D. W.; Mautner, M. N. Biochimica et
Biophysica Acta, Biomembranes 2002, 1559, 1.

(62) Forster, T. Ann. Physik. 1948, 2, 55.

(63) Jones, G. M.; Wofsy, C.; Aurell, C.; Sklar, L. A. Biophysical Journal
1999, 76,517.

(64) Keller, L.; Hussey, D. M.; Fayer, M. D. Journal of Physical
Chemistry 1 996, 100, 10257.

137

 

(65) Merta, J.; Garamus, V. M.; Kuklin, A. I.; Willumeit, R.; Stenius, P.
Langmuir 2000, 16, 10061.

(66) Klein, U. K. A.; Haar, H. P. Chemical Physics Letters 1979, 63, 40.
(67) Labhart, H.; Pantke, E. R. Chemical Physics Letters 1973, 23,482.

(68) Youngren, G. K.; Acrivos, A. Journal of Chemical Physics 1975, 63,
3846.

(69) Lipari, G.; Szabo, A. Biophysical Joumal 1980, 30,489.

(70) Debye, P. Polar Molecules. In Chemical Catalog Co. New York,
1929.

(71) Goldie, S. N.; Blanchard, G. J. Joumal of Physical Chemistry A
1999, 103,999.

(72) Jiang, Y.; Blanchard, G. J. Journal of Physical Chemistry 1 994, 98,
6436.

(73) Jiang, Y.; Blanchard, G. J. Journal of Physical Chemistry 1995, 99,
7904.

(74) Zinsli, P. E. Chemical Physics 1977, 20, 299.

(75) Einstein, A. Annalen der Physik (Weinheim, Germany) 1905, 17,
549.

138

 

Chapter 6

CONCLUSIONS AND FUTURE WORK

In this dissertation, we have studied the fluorescence lifetime and rotation
diffusion dynamics of perylene in micellar and vesicular environments. For the
micellar systems, we have concluded that the electric double layer plays a
substantial role in mediating interfacial interactions. For the micelle and vesicle
systems, we have concluded that there are differences in the structures. To
better understand the micellar and vesicular interfacial region, more studies
regarding the specific differences between the two structures are needed.

The work performed in chapter 2 of this dissertation shows that the
counterions present in the micellar solutions do indeed effect the environment of
the micellar interior. The reason for the non-monotonic variations in the
microviscosities of the micellar interiors was inconclusive with the available data.
in chapter 3, we attempt to explain the non-monotonic variations by the presence
of a cosolvent in the micelle solutions. The data we present there collectively
point to the existence of microscopic heterogeneity in these binary solvent
systems. Altering the counterions of the anionic and cationic micellar systems or
performing similar experiments on a nonionic surfactant system would provide
additional insight into the role of the double layer in mediating solvation.

In chapters 4 and 5 of this dissertation, the differences between micelle
and vesicle systems were examined and we determined that the organization of

the two structures was notably different. More specifically, the micelle systems

139

were found to have a more rigid structure than their bilayer or vesicle
counterparts. We believe that this is due to the difference in the size and the radii
of curvature of the two structures. We also found that increasing the acceptor
concentration allows multiple acceptors to form complexes with the vesicle
surface but not with the micelle surface, a fact which we contribute to the
different charges present on the micelle and vesicle surfaces. Future work with
this research would entail using the information gained here about micellar and
vesicular structures and applying similar studies to phospholipid vesicular
systems. Another avenue for scientific exploration would be trying similar studies
on langmuir—blodgett films, to probe the differences between curved and flat

StI'UCtU 1'68.

140

 

5

ililiilliiiifllililisljliji1