DIFFUSIONAL MOTION AS A GAUGE OF INTERFACIAL FLUIDITY AND ADHESION
OF SUPPORTED MODEL MEMBRANE FILMS
By
Stephen M Baumler

A DISSERTATION
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
Chemistry – Doctor of Philosophy
2017

ABSTRACT
DIFFUSIONAL MOTION AS A GAUGE OF INTERFACIAL FLUIDITY AND ADHESION
OF SUPPORTED MODEL MEMBRANE FILMS
By
Stephen M Baumler
The plasma membrane plays an important role in cellular processes. Not only does the
membrane act as a barrier to the outside world, compartmentalizing cell functions by providing a
two-dimensional solvent, but directly, by changing protein conformation and function, and
indirectly, by changes in diffusional properties of the membrane through changes in chemical
composition. The utility of model membrane systems in chemical sensing is predicated upon
housing transmembrane proteins in their active conformation(s). Therefore, an understanding of
the role the supporting surface has on mimicking the physical properties of plasma membranes is
necessary in the creation of biomimetic films that maintain the functional properties of the
imbedded proteins.
Measuring diffusional motions via time-resolved picosecond laser spectroscopies and
fluorescence microscopies, the fluidity and interfacial adhesion strength at an organic/inorganic
interface of both monolayer thin films and supported lipid bilayers has been examined. Films have
been prepared using both Langmuir-Blodgett deposition and small unilamellar vesicle (SUV)
deposition. Both physical and chemical interactions were found to play an important role in
mediating the dynamical properties exhibited by these films. First, the addition of an aqueous
adlayer to lipid films produced using SUV deposition was found to significantly disrupt the
organizational order within the alkyl chain region.

Second, by combining rotational and

translational diffusion measurements from a tethered fluorescent probe molecule, compositional
variations within alkylphosphonic acid Langmuir films induced varying degrees of adhesion and

fluidity. Lastly, the extent to which the fluidity of a supported lipid film is mediated by the ionic
interactions between head-group and supporting surface vs. that of the lipid-lipid tail-group
interactions was examined and shown to be system dependent.
This research has led to an adaptable methodology based on the use of rotational and
translational diffusional data to quantitate the strength of supported films over a range of length
scales (sub-diffraction limit – to – hundreds of microns). A novel family of interfaces that can be
physically bound to a surface and at the same time have controlled fluidity depending upon a range
of chemical conditions of the underlying inorganic support has been developed.

These

developments move closer to providing an understanding of the impact a supporting surface plays
in mediating model membrane dynamical properties and function.

Copyright by
STEPHEN M BAUMLER
2017

To Mom, Dad, Kathy, and Ronny.

v

ACKNOWLEDGMENTS

I would like to sincerely thank everyone who has contributed to my personal and
professional development over the past few years. First and foremost, I would like to thank my
research advisor Professor Gary J. Blanchard. I am thankful for the freedom you gave me to pursue
different projects, and the many helpful suggestions when they failed. Without your mentorship,
I would not be the scientist I am today.
I also want to express my gratitude to my committee members: Professors Tom Hamann,
Jim McCusker, and Greg Swain for their guidance and insightful suggestions. Thanks to Dr. Kathy
Severin, and Professors Merlin Bruening and Greg Swain for their mentorship in teaching the
many semesters of the analytical and physical chemistry lab sequence. I am thankful to the
Michigan State University Chemistry Department, and United States Department of Education
(GAANN Fellowship) for funding this project and allowing me to perform this research.
Many thanks to my colleagues and lab mates for the fun times inside and outside of lab. I
would like to specifically thank Dr. Omid Zandi and Josh Baillargeon for their friendship,
camaraderie, and enjoyment from countless hours of fruitful science discussions at the local
watering holes. I would also like to thank past and present Blanchard group members for truly
making graduate school a wonderful experience.
I could not be more thankful for the constant support from my parents, brother, family and
friends. Specifically, I would like to thank my parents for their love, devotion and unwavering
support throughout my education. Finally, I would like to thank Emily. Thank you for your
continued optimism, putting up with my long hours at work, support and encouragement. I am
truly grateful and look forward to many future adventures.

vi

TABLE OF CONTENTS

LIST OF TABLES ......................................................................................................................... ix
LIST OF FIGURES .........................................................................................................................x
CHAPTER 1: INTRODUCTION ...................................................................................................1
1.1 Motivation ............................................................................................................................2
1.2 Lipid Bilayer Membranes ....................................................................................................2
1.3 Model Membrane Systems ..................................................................................................4
1.4 Objectives.............................................................................................................................8
REFERENCES ........................................................................................................................10
CHAPTER 2: EXPERIMENTAL TECHNIQUES.......................................................................15
2.1 Instrumental Methods for the Examination of Lipid Dynamics ........................................16
2.2 Evaluating Film Morphology with Brewster Angle Microscopy (BAM)..........................18
2.3 Measuring Diffusion: Time-Resolved Fluorescence Spectroscopy ...................................21
REFERENCES ........................................................................................................................35
CHAPTER 3: LIPID ADLAYER ORGANIZATION MEDIATED BY A LIQUID
OVERLAYER ...............................................................................................................................40
3.1 Abstract ..............................................................................................................................41
3.2 Introduction ........................................................................................................................41
3.3 Experiment .........................................................................................................................43
3.4 Results and Discussion.......................................................................................................49
3.5 Conclusions ........................................................................................................................60
REFERENCES ........................................................................................................................62
CHAPTER 4: USING DIFFUSIONAL MOTION TO GAUGE INTERFACIAL ADHESION
OF SUPPORTED ALKYL PHOSPHONATE MONOLAYERS .................................................67
4.1 Abstract ..............................................................................................................................68
4.2 Introduction ........................................................................................................................68
4.3 Experimental ......................................................................................................................70
4.4 Results and Discussion.......................................................................................................77
4.5 Conclusions ........................................................................................................................94
REFERENCES ........................................................................................................................96
CHAPTER 5: THE INFLUENCE OF METAL IONS ON THE DYNAMICS OF SUPPORTED
PHOSPHOLIPID LANGMUIR FILMS ......................................................................................102
5.1 Abstract ............................................................................................................................103
5.2 Introduction ......................................................................................................................103
5.3 Experimental ....................................................................................................................105
5.4 Results and Discussion.....................................................................................................109
5.4 Conclusions ......................................................................................................................122
APPENDIX ............................................................................................................................123
REFERENCES ......................................................................................................................132

vii

CHAPTER 6: CONCLUSIONS AND FUTURE DIRECTIONS ..............................................137
6.1 Conclusions and Future Directions ..................................................................................138
REFERENCES ......................................................................................................................144

viii

LIST OF TABLES

Table 3-1: Reorientation time (𝜏𝜏OR), Rotational Diffusion Constant (D), and Calculated Local
Viscosity (Ρ) determined by Local Environment for DMPC Bound to a Modified Silica
Substrate.................................................................................................................................. 52
Table 3-2: Adlayer ellipsometric thickness and electrochemical coverage. ................................. 59
Table 4-1: Translational diffusion constants (DT), Mobile fraction %, and Recovery speed (τ1/2)
determined for the 5 mol% and 10 mol% substrates from FRAP measurements. .................. 85
Table 4-2: Reorientation time (𝜏𝜏HR), rotational diffusion constant (DROT), and rotational wobbling
constant (DW) determined by the local Environment of BLPA in a Ba-ODPA LB film. ....... 88
Table 4-3: Calculated Diffusion constant (DT), Unbound free fraction (Afree), Dissociation
equilibrium constant (Keq), and Free energy for dissociation (ΔGdis) for the BLPA in a BaODPA LB film. ....................................................................................................................... 93
Table 5-1: Relative metal ion surface coverage for the phosphorylated FTO and glass surfaces are
reported versus phosphorus (P2p), tin (Sn3d5/2), and silica (Si2p) peak areas. .................. 113
Table A5-1: Dissociation constant (koff) data acquired by fitting FRAP curves with the reactiondominant model. ................................................................................................................... 128
Table A5-2: Diffusion coefficient data acquired by fitting FRAP curves with the full reactiondiffusion model. .................................................................................................................... 128
Table A5-3: One-way ANOVA test among all surfaces. ........................................................... 130
Table A5-4: One-way ANOVA test among all metal surfaces................................................... 130
Table A5-5: One-way ANOVA test among alkaline metal surfaces .......................................... 130
Table A5-6: One-way ANOVA test among heavy metal surfaces. ............................................ 130

ix

LIST OF FIGURES

Figure 1-1: The amphiphilic nature of a lipid molecule is shown as a combination of the
hydrophilic headgroup and hydrophobic tail (left). The amphiphilic nature of lipids gives rise
to the self-assembly of lipid bilayers in the presence of an aqueous environment (right)........ 3
Figure 1-2: Molecular structure for a typical lipid is shown in (a.) Both R2 and R3 consist of alkyl
chains of varying lengths and degrees of saturation. R1 corresponds to the esterified side group
giving the lipid a unique charge at different pH values, commonly named (b.) phosphatidic
acid, (c.) phosphatidylglycerol, (d.) phosphatidylinositol, (e.) phosphatidylserine, (f.)
phosphatidylethanolamine, and (g.) phosphatidylcholine. ....................................................... 4
Figure 2-1: Description of the application of Brewster's law for the visualization of the formation
dynamics of Langmuir monolayers at an air/water interface.................................................. 19
Figure 2-2: Schematic of the Brewster Angle Microscope (BAM) system. ................................. 20
Figure 2-3: Schematic for the Fluorescence Anisotropy Decay Imaging (FADI) instrument. ..... 26
Figure 2-4: Visual representation of the photon counting scheme for time correlated single photon
counting (TCSPC)................................................................................................................... 27
Figure 2-5: Reference, signal, and normalized response FRAP curve of perylene in a solid
supported DMPC bilayer. ....................................................................................................... 29
Figure 2-6: Functional form for FRAP curves based on the Soumpasis model for various diffusion
times “τD” (s) (left) and spot sizes (μm) (right). The best fit for a recovery curve of perylene in
a solid supported DMPC bilayer is shown with the residuals below. ..................................... 30
Figure 2-7: Schematic for the Fluorescence Recovery After Photobleaching (FRAP) instrument.
................................................................................................................................................. 33
Figure 3-1: Chemical Structures of 1,2-dimyristoyl-sn-phosphatidylcholine (DMPC) and Perylene.
................................................................................................................................................. 44
Figure 3-2: Steady state excitation (S1 ← S0, solid line) and emission (S0 ← S1, dashed line) of
perylene in DMPC vesicles prior to formation of the lipid monolayer. ................................. 45
Figure 3-3: Idealized diagram of the prepared supported lipid layer. The films are composed of
the (a) gold or silicon support, (b) phosphorylated APDMES (c) zirconation layer, and (d)
DMPC layer. ........................................................................................................................... 47
Figure 3-4: Induced orientational anisotropy decay data for perylene in DMPC monolayer on SiOx
substrates, (a.) exposed to air and (b.) exposed to an aqueous overlayer. The solid lines are the
best fit, single exponential decay function. Insets represent residuals of fit........................... 51

x

Figure 3-5: Cyclic voltammograms of a blank Au surface (─), a C12 thiol monolayer (---) and a
DMPC adlayer bound to Au (⋯⋯). All CVs were taken in 0.1M KCl (aq) solution at a scan rate
of 100 mV/s............................................................................................................................. 55
Figure 3-6: AC impedance for a blank Au surface (●), a C12 thiol SAM on Au (■), and a DMPC
adlayer bound to Au (▲). Inset: Equivalent circuit used to extract Rct values from these data:
Ru = uncompensated resistance, Rct = charge transfer resistance, Cdl = double layer capacitance.
................................................................................................................................................. 57
Figure 4-1: Chemical structures of the components used in the formation of the LB film.
Octadecylphosphonic acid (ODPA), and 1-(12-[4-(dipyrrometheneboron difluoride) butanoyl]
amino) dodecanoyl-2-hydroxy-sn-glycero-3-phosphate (ammonium salt) (BLPA). ............. 71
Figure 4-2: Idealized diagram of the prepared ODPA/BLPA Langmuir film. The films are
composed of the (a) phosphorylated silica support, (b) barium ion layer, and (d) ODPA/BLPA
layer......................................................................................................................................... 73
Figure 4-3: Above: 𝜋𝜋/A-isotherm of ODPA on an aqueous subphase (red solid) and with a 5 mM
Ba2+ subphase (blue dash). 𝜋𝜋 /A-isotherm of BLPA on an aqueous subphase (red dot) and with
a 5 mM Ba2+ subphase (blue dot). The temperature for the subphase was 23 °C and pH = 5.4.
The monolayers were compressed at a rate of 0.8 Å2/min. Orange circles represent pressures
at which BAM images (A–F) were taken. Below: Images A-C are of ODPA on an aqueous
subphase. Images D-F are of ODPA on an aqueous subphase containing 5 mM BaCl2. Scale
bars represent 100 Îźm. ............................................................................................................ 79
Figure 4-4: Above: 𝜋𝜋/A-isotherm of the two component monolayers containing ODPA and BLPA
on an aqueous subphase with a Ba2+ concentration of 5 mM, and 0 mol% BLPA (red solid), 5
mol% (blue dash), and 10 mol% (green dot). The temperature for the subphase was 23 °C and
pH = 5.4. The monolayers were compressed at a rate of 0.8 Å2/min. Orange circles represent
pressures at which BAM images (A-F) were taken. Below: Images A-C contain ODPA and 5
mol% BLPA, images D-F contain ODPA and 10 mol% BLPA. Images at 0 mol% BLPA are
presented in Fig. 4-3 (D-F). Scale bars represent 100 Îźm. .................................................... 81
Figure 4-5: π/A-isotherm of the two component monolayers containing ODPA and BLPA on an
aqueous subphase with a Ba2+ concentration of 5 mM, and 0 mol% BLPA (100% ODPA) (red
dash), 5 mol% (blue dot), and 10 mol% (blue dash-dot) dissolved in THF when applied to
trough. The application of ODPA when dissolved in chloroform at 0.2 mg/mL and added under
similar conditions is shown (red solid). .................................................................................. 82
Figure 4-6: (A.) Emission spectra of the two component LB films. BLPA-5 (blue solid), and
BLPA-10 (green dash). Absorbance and emission spectra of BLPA in THF (black dot). All
spectra were normalized to unity. Schematic representation of a structural description of Hand J-aggregates of the BODIPY chromophore (B.) Steady state fluorescence images of
BLPA-5 are shown in (C.) and BLPA-10 in (D.). Scale bars represent 500 Îźm. .................. 84
Figure 4-7: FRAP recovery of the BLPA chromophore in (A.) BLPA-5 (red) and (B.) BLPA-10
(blue). The green lines represent best-fit lines using the Ellenberg equation with residuals

xi

plotted below. (C.) Comparison between the BLPA-5 and BLPA-10 curves is shown. All
FRAP curves were normalized prior to fit. ............................................................................. 86
Figure 4-8: Typical experimental anisotropy decay functions for the BLPA-5 (A.) and BLPA-10
(D.) LB films with residuals of best fit inset are shown. Fluorescence transients were fit to
determine the fluorescence lifetime shown in (B., BLPA-5) and (E., BLPA-10). The colors
shown represent the lifetime value at a given pixel, with blue being short lifetimes and red
being longer lifetimes. Fluorescence anisotropy decay time constant images are shown in (C.,
BLPA-5) and (F. BLPA-10). The colors shown are set by the imaging software, with blue
being short decay time constants and red being longer decay time constants. All images were
acquired using a 10x microscope objective. Scale bars represent 500 Îźm. ............................ 87
Figure 4-9: Schematic of equilibrium system under consideration. ............................................. 89
Figure 5-1: Chemical structures of 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC) and 1(12-[(7-nitro-2-1,3-benzoxadiazol-4-yl)amino]dodecanoyl)-2-hydroxy-sn-glycero-3phosphocholine (NBD-LPC). ............................................................................................... 106
Figure 5-2: Surface pressure / mean molecular area-isotherm of the DMPC/NBD-LPC monolayers
formed on subphases containing buffer (red solid), and 1 mM MgCl2 (blue dash), CaCl2 (green
dot), or BaCl2 (purple dash-dot). Inset: Expanded region showing mean molecular area at
dipping pressure. ................................................................................................................... 110
Figure 5-3: Surface pressure / mean molecular area-isotherm of the DMPC/NBD-LPC monolayers
formed on subphases containing buffer (red solid), and 1 mM NiCl2 (blue dash), ZnCl2 (green
dot), CdCl2 (purple dash-dot), or ZrOCl2 (orange dash). Inset: Expanded region showing mean
molecular area at dipping pressure. ...................................................................................... 111
Figure 5-4: Interactions between the metal ions and lipid monolayer are shown as the mean lipid
area vs. Metal ion van der Waal radius. Error bars are calculated by extrapolation of a line
tangent to the isotherm between 20-30 mN/m down to 0 mN/m.......................................... 112
Figure 5-5: Idealized diagram of the prepared supported lipid layer. The films are comprised of
the phosphorylated glass support (a.), metal ion and aqueous under-layer (b.), and
DMPC/NBD-LPC film (c.) with buffer adlayer (d.)............................................................. 114
Figure 5-6: Normalized FRAP recovery curve from one SiOx surface fitted to the full reactiondiffusion model. Residuals are plotted below. Inset: Average of all trials on the SiOx (red
circles) and phosphorylated surface (blue squares) are plotted with error (standard deviation)
represented as shading. ......................................................................................................... 115
Figure 5-7: Bleaching spot area vs. lipid dissociation rate (koff) determined by the reaction
dominant model. Error bars represent 95% confidence interval........................................... 117
Figure 5-8: Translational diffusion coefficients acquired from fitting the full reaction diffusion
model for each prepared surface. Error bars represent 95% confidence interval. ............... 118

xii

Figure 5-9: Log of formation constants for methylphosphonic acid to each metal ion vs.
translational diffusion coefficient. Error bars represent 95% confidence interval. ............. 120
Figure 5-10: Idealized representation of the energies required for the NBD-LPC to undergo
translational diffusion in a DMPC layer. The translational diffusion is governed by both an
activation energy from the metal phosphonate-lipid interaction (EMP-L) and activation energy
from the lipid-lipid van der Waals dispersion interactions (EL-L). The free energy for
translational diffusion (ΔGDt) is therefore governed by the contributions from both metal
phosphonate-lipid (ΔGMP-L) and lipid-lipid interactions (ΔGL-L). ........................................ 121
Figure A5-1: XPS survey spectra of DMPC (red), Mg2+ (blue), and Ba2+ (green) are shown…125
Figure A5-2: (left). Fit of the XPS spectral data for a DMPC/NBD-LPC film containing Ba2+ is
shown. The Ba3d5/2 peak is fit to a Gaussian-Lorentzian curve after Shirley background
subtraction... .......................................................................................................................... 125
Figure A5-3: (right). Representative convoluted fits for the overlapping Sn4s and P2p peaks on
the Ba2+/DMPC/NBD-LPC film. .......................................................................................... 125
Figure A5-4: Box plot for the fitted diffusion coefficients of all trials on each surface. The red line
represents Median, whiskers represent maximum and minimum values, and red asterisks are
considered outliers. Notches in the box plot represent 95% confidence intervals for each
surface. .................................................................................................................................. 129

xiii

CHAPTER 1: INTRODUCTION

1

1.1

Motivation
Since the introduction of the fluid mosaic model, the primary role of lipids has been thought

to be that of a barrier to the outside world; compartmentalizing cell functions by providing a twodimensional solvent for the myriad of proteins performing various tasks.1 Continued research
suggests that lipids play a much more important role. The plasma membrane plays an important
role in cellular processes both directly, by mediating protein conformation and function, and
indirectly, via changes in diffusional properties of the membrane.2-3 The study of transmembrane
processes at the molecular level, such as transport of ions and molecules, cell signaling and
adhesion, have relied upon the formation of cell membrane model systems such as supported lipid
monolayers and bilayers (SLBs) and black lipid membranes.4 While these model systems create
a biomimetic environment for the study of cellular structure-function relationships, examination
of a means to control the dynamical properties of such systems has been limited. The aim of this
dissertation is to better understand ways to characterize and control the dynamical properties of
the supported films, with the long-term goal of utilizing this knowledge to create and control the
functional properties of a biomimetic bilayer structure.

1.2

Lipid Bilayer Membranes
Membranes are ubiquitous in all living things and are a complex barrier to the outside

world. The membrane not only serves as a means to protect a cell from the external environment,
but is the critical interface at which chemical transport is regulated and cellular signal transmission
occurs.

At the most basic level, plasma membranes consist of a bilayer of amphiphilic

biomolecules consisting of lipids (phospholipids and cholesterol), biopolymers (peptides and
proteins), and carbohydrates.

2

Figure 1-1: The amphiphilic nature of a lipid molecule is shown as a combination of the
hydrophilic headgroup and hydrophobic tail (left). The amphiphilic nature of lipids gives
rise to the self-assembly of lipid bilayers in the presence of an aqueous environment
(right).
Lipids are amphiphilic molecules containing both a hydrophilic region (headgroup) and
hydrophobic region (tail) (Fig. 1-1). In the presence of water, the amphiphilic nature of the
phospholipid results in their self-assembly into a bilayer structure with the headgroup in contact
with water, and the tails facing each other.
Native plasma membranes consist of thousands of unique chemical species, with over 500
unique lipids in the human plasma membrane alone.5 Lipids are commonly sorted by both their
headgroup type, and tail length and degree of saturation. The five most common lipid headgroups
are shown in figure 1-2. The different headgroups are a result of the esterification of a single
oxygen on the phosphonic acid group attached to the phospholipid (Fig. 1-2, a.). The headgroup
is important in defining both the overall charge at the lipid-water interface as well as contributing
to lipid-lipid interactions through electrostatic interactions or hydrogen bonding. For example, at
pH = 7, headgroups containing hydroxyl groups, with the exception of phosphatidylserine, will

3

Figure 1-2: Molecular structure for a typical lipid is shown in (a.) Both R2 and R3 consist
of alkyl chains of varying lengths and degrees of saturation. R1 corresponds to the
esterified side group giving the lipid a unique charge at different pH values, commonly
named (b.) phosphatidic acid, (c.) phosphatidylglycerol, (d.) phosphatidylinositol, (e.)
phosphatidylserine, (f.) phosphatidylethanolamine, and (g.) phosphatidylcholine.
add zero charge to the lipid headgroup, giving a net (-1) charge to the lipid. In comparison, both
ethanolamine and choline contribute a (+1) charge to the headgroup, giving the lipid zwitterionic
character or a net neutral charge.6
Over the life of a typical cell,
morphological changes occur.7-8

diverse and dynamic chemical compositional and

Steinman, et. al. have shown that the plasma membrane

experiences a complete turnover or recycling of lipids over the course of an hour.9 As a result of
the constant turnover of lipids, the formation of membrane patches or domains occurs lasting tens
of seconds, but disperse soon after.10 Developing a biomimetic platform, whether for the study of
transmembrane processes or utilization as a biosensor, that maintains the fully functional form of
the proteins without the full morphological and compositional complexity of the native
environment, remains a challenge.

1.3

Model Membrane Systems
There are many different types of model membrane systems, but are commonly grouped

as either supported or unsupported model membrane systems. Below, a brief review of a few

4

relevant methods for preparing model membrane systems, their limitations and applications, with
the focus on solid supported membranes is examined.

Unsupported Lipid Membranes. Unsupported lipid membranes are relatively easy to
fabricate and maintain similar dynamic properties to native plasma membranes. The preservation
of the natural properties of plasma membranes in unsupported lipid bilayers allows for the
embedding of transmembrane proteins that maintain their functional form. Unsupported lipid
membranes are limited in their stability (typically hours), availability of analytical techniques to
characterize their properties, and need for a solid support to act as a transducer for many biotechnological applications (biosensors, separations, self-healing and tribological surfaces).11

Black Lipid Membranes (BLMs). Early work on the formation and examination of lipid
membrane models involved using a Langmuir trough to self-assemble lipids at the air-water
interface and transfer to a small aperture (typically < 1mm diameter) contained on a thin film of
Teflon.12-13 The membranes were termed “black” due to the observation that under reflected light,
the birefringent quality of the pinhole region became non-reflecting over time.14 Fabrication of
BLMs leaves both outer in inner leaflets available to interrogation. As a result of the easy access
to both interfaces, BLMs have found wide use in the investigation of ion-channel activity and
membrane permeability studies.15 Despite recent efforts to decrease the aperture size to submicrometer diameters for increased structural robustness, these systems have yet to exhibit longterm stability.16-18

5

Liposomes (Lipid Vesicles). Liposomes are spherically structured lipid bilayers containing
an internal volume of an aqueous phase and are typically classified by their size and degree of
lamella.19-22 Multilamellar vesicles (MLVs) are vesicles containing more than a singular bilayer.
Because of their ill-defined geometry, and lack of structural uniformity, MLVs are rarely used for
analytical study. MLVs form readily by self-assembly when dried lipids are introduced to an
aqueous phase. They are often an observed initial structure in the preparation small unilamellar
vesicles (SUVs, diameter < 100 nm), large unilamellar vesicles (LUVs, d > 100 nm), and giant
unilamellar vesicles (GUVs, d > 10 Îźm). Recently, lipid vesicles have been extensively studied
and found widespread use as liposomal drug delivery systems.23 Despite these successes, vesicles
remain less than ideal candidates for studying protein-lipid interfaces and for use in
biotechnological applications. The curvature induced by formation of the vesicles is much higher
compared to the curvature observed in cell membranes, resulting in packing constraints different
than the native plasma membrane.24 Furthermore, polydispersity in vesicle size gives rise to
curvature induced differential responses in the physical properties of the lipid-lipid or lipid-protein
interface limiting the ability to precisely measure a given property.25-26

Supported Lipid Membranes. In the mid-1980s Tamm and McConnell first showed the
viability of supporting phospholipid bilayers at a solid interface.27 The method presents a one-step
procedure for creating solid supported model membranes containing different lipid mixtures.
Since then, there have been extensive studies on the use of supported lipid bilayers (SLBs) as a
tool to understand lipid domain formation, and protein membrane interactions.28-31

Recent

advancements combining both quartz crystal microbalance with dissipation monitoring (QCM-D)

6

and atomic force microscopy (AFM) have led to an understanding of the mechanisms involved in
the formation of these SLB systems.32-33
In general, there are two methods commonly used for the formation of SLBs. The first
method involves the deposition of small unilamellar vesicles (SUVs) onto the supporting surface.
Recent studies have examined the extent to which the composition of the underlying support
influences the formation and adhesion of supported lipid bilayers prepared by the fusion deposition
of small unilamellar vesicles (SUVs). The identity of the underlying support has a marked effect
on both the rupture and spreading processes of the SUVs.34-38 The second method utilizes the
Langmuir-Blodgett or Langmuir-Schaffer technique to prepare monomolecular or multilayer
films.

39-41

The use of the Langmuir-Blodgett technique provides a controlled means for the

formation of these SLBs in comparison to the SUV approach. Because these films are built one
molecular layer at a time, the bilayer can be built asymmetrically, a property of natural membranes
that is not accessible using the SUV approach.42-43
SLBs can be categorized into two main types based upon the supporting interface. The
simplest type consists of a mono- or bilayer on a solid support. Formation of SLBs on a solid
support has seen a variety of supporting materials such as silica, glass, mica, various metal oxides
such as TiO2 and SrTiO3, and more recently inorganic nanoparticles.34-38 The formation of such
films is relatively simple and straightforward provided the proper pH and ionic strength during
preparation.44 The second type of supported lipid membrane, commonly referred to as hybrid
bilayer membranes (HBMs)45-47 contain an added organic (self-assembled monolayer) or
biological (protein or peptide) cushioning between the supporting surface and lipid layer.48-50
Often, these HBMs are formed at a metallic interface lending themselves amenable for use as

7

electrochemical biosensors.51 Unfortunately, these systems are often plagued with poor stability
in air and exhibit slowed dynamical properties relative to native membranes.30, 52

1.4

Objectives
Utilizing the formation and adhesion of SUVs, the Blanchard group has shown previously

that robust air-stable lipid layers can be formed through the aid of zirconium-phosphate (ZP)
chemistry and is dependent upon the lipid head-group used in self-assembly process.53 The lipid
adlayers formed by this approach were found to be structurally complete and nominally
homogeneous monolayers. Later studies examined how the underlying metal ion identity mediated
the lipid-surface interactions during the formation of the lipid-metal-phosphonate films.54 The
formation of these lipid films through the lipid-metal coordination process was found to rely upon
the use of metal ions with high charge and small ionic radius. These studies showed the possibility
of the formation of air stable lipid layers, and how variations in the metal ions used changed the
permeability and thickness. The dynamical properties of these films were unknown.
Using the foundational work on the formation of supported and air-stable lipid layers, the
work in this dissertation has focused on understanding how the physical interactions occurring at
the lipid-aqueous, lipid-support, and lipid-lipid interfaces give rise to the observed dynamical
properties for a given system. We used Time Correlated Single Photon Counting (TCSPC)
Microscopy and Fluorescence Recovery After Photobleaching (FRAP) as measurement techniques
to examine the rotational and translational behavior of probe molecules in a variety of supported
lipid systems, and provide a detailed description of the measurement science in Chapter 2. In
Chapter 3, we first examined the role of an aqueous overlayer on the dynamics of a perylene probe
embedded within a covalently bound lipid monolayer using TCSPC and characterized the

8

permeability of the interface using electrochemical potential and impedance measurements. The
combination of the rotational and translational diffusion properties of an alkylphosphonic acid
Langmuir film is used to understand the molecular-micro scale heterogeneity, fluidity, and
adhesion strength of the film at the supporting interface in Chapter 4. In Chapter 5, using FRAP
and x-ray photoelectron spectroscopy (XPS), we examined the role that different alkaline earth
and transition metals play in modulating the fluidity of a supported lipid film. The combination
of optical techniques comparing different physical properties on different time-scales has opened
the possibility to study the complex dynamic properties inherent to both supported lipid
membranes and their natural plasma membrane counterpart.

The ability to measure these

properties allows for both the creation of films with modulated fluidity, and the ability to
understand the dynamic conditions necessary to maintain fully functional transmembrane proteins
within a model system.

9

REFERENCES

10

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Crane, J. M.; Kiessling, V.; Tamm, L. K., Measuring lipid asymmetry in planar supported
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Kiessling, V.; Wan, C.; Tamm, L. K., Domain coupling in asymmetric lipid bilayers. BbaBiomembranes 2009, 1788 (1), 64-71.

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Plant, A. L., Supported hybrid bilayer membranes as rugged cell membrane mimics.
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Plant, A. L.; Tarlov, M., Self-Assembled Monolayers as Substrates for Planar Model
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Albertorio, F.; Diaz, A. J.; Yang, T. L.; Chapa, V. A.; Kataoka, S.; Castellana, E. T.;
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Matyszewska, D.; Bilewicz, R.; Su, Z. F.; Abbasi, F.; Leitch, J. J.; Lipkowski, J., PMIRRAS Studies of DMPC Bilayers Supported on Au(111) Electrodes Modified with
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14

CHAPTER 2: EXPERIMENTAL TECHNIQUES

15

2.1

Instrumental Methods for the Examination of Lipid Dynamics
The organization of lipids within a bilayer structure creates a dynamic environment on a

variety of length and timescales. The dynamic properties of the membrane are determined by each
lipid component contributing unique conformational, translational, and molecular rotation
dynamics within the membrane as a whole. In addition, the possibility of collective motions of
the entire membrane or individual subgroups can exist. For example, the dynamics of lipids exhibit
vastly different conformational, translation, and rotation dynamics in comparison to their protein
and peptide counterparts. The examination of even simple model lipid membranes remains a
challenge because the unique dynamical properties cannot be simply probed by a single physical
parameter. While not an exhaustive list, a variety of magnetic, optical, acoustic and scanning
probe methods exist for the examination of different membrane properties. The utility and
limitations of the most common and widely used methods for study of membrane dynamics will
be discussed briefly below.

Magnetic Resonance Methods. Both electron paramagnetic resonance (EPR) and nuclear
magnetic resonance (NMR) have been used to examine a variety of dynamical properties of
membranes. Typically, these techniques are used to examine the conformational order of the lipid
alkyl chain region based on spectrum splitting of a site-directed spin label (EPR) or quadrupole
splitting of deuterated lipid molecules (NMR).1-2 The use of site-directed spin labeling (SDSL), a
now common EPR technique, has proved useful to examine the lipid-protein interface because the
sensitivity of the measurement technique is matched to the rotational dynamics of lipids
(nanoseconds).3-4 Using deuterium labeled lipid alkyl chains of specific lipid types, researchers
have been successfully using 2H solid state NMR to examine gauche defects and ordering of the

16

tail region at the lipid-lipid and lipid-protein interface.5-6

While both techniques provide

information on the molecular details of the tail orientation and order, they are limited by a variety
of factors. The measurements are an average response from the model under study, so detection
of heterogeneity on a scale larger than the molecular scale is difficult.

Optical Spectroscopy Methods. Linear and non-linear optical spectroscopy techniques
have been successfully used to examine a variety of dynamical properties of lipid membranes.
Similar to EPR and NMR methods, different forms of IR spectroscopy have been employed to
examine the structural order of the long lipid alkyl chain groups in the bilayer. Polarization
modulation infrared reflection absorption spectroscopy (PM-IRRAS) is commonly coupled to a
Langmuir trough allowing for the study of changes in conformation of both lipid alkyl chain and
headgroups of a single monolayer at the air-water interface.7-8 Nonlinear methods such as second
harmonic generation (SHG) and sum frequency generation (SFG) have been successfully used to
examine the dynamics of lipid diffusion perpendicular to the bilayer plane (lipid flip-flop or
translocation) as well as changes in the conformational order of the lipid tail groups.9-13 The most
commonly employed methods for the study of translational and rotational diffusion in membranes
are time resolved fluorescence techniques. Fluorescence correlation spectroscopy (FCS) has been
in use since the early 1970s to examine chemical rate constants and diffusion coefficients of
biomolecules in solution, however it was not until the mid-1990s that the technique overcame
previous limitations of large background signals and low signal count rates.14-17

The low

concentrations of chromophores necessary for the technique are beneficial due to decreased
morphological disruption of the model system. Unfortunately, FCS is also limited by the need for

17

low concentrations in that the inherent heterogeneity can provide poor precision in observed values
or go unobserved altogether.18

Scanning Probe and Acoustic Methods. Both quartz crystal microbalance with dissipation
monitoring (QCM-D) and atomic force microscopy (AFM) have contributed greatly to the
understanding of the mechanism by which SUVs adhere and form stable supported bilayer
systems.19-22 Both methods provide unique insight into the fluid-like properties of model cell
membranes, however they are limited in their ability to spatially resolve large-scale (100 μm – to
– cm) heterogeneities within the supported lipid bilayers. QCM-D measures the average response
from changes in frequency and dissipation energy over the size of the sample, typically 1-2 cm
diameter disk, and therefore has very limited spatial resolution. The spatial resolution of AFM is
unparalleled relative to the other techniques discussed within this dissertation, however the total
observed area makes observations of macroscale compositional changes difficult.

2.2

Evaluating Film Morphology with Brewster Angle Microscopy (BAM)
Analytical methods for the direct observation of large scale (Îźm-to-cm) morphological

features are often limited to the use of dyes or stains and an optical microscope. During preparation
of Langmuir-Blodgett films, the addition of dyes or stains could potentially disrupt the
morphological features of a monolayer film, even at relatively low concentrations. Additionally,
the resolution is often limited by the depth of focus of the microscope objective at a given
magnification. Brewster Angle Microscopy (BAM) was first introduced in 1991 and is currently
the most informative method for the examination of relatively largescale (Îźm-to-cm) changes to
Langmuir films under a variety of conditions.23

18

Sir David Brewster first observed when “a pencil of light is incident upon the separating
surface of two media having different indices of refraction m m’, it will be polarised at an angle
whose tangent is equal to the quotient of the greater index of refraction divided by the lesser, or
m/m’.”24 This observation is now commonly known as Brewster’s law (eqn. 2-1), where θB is
Brewster’s angle, or the angle at which maximum polarization occurs, n1 and n2 are the indices of
refraction of the two materials.
tan θ B =

n1
n2

2-1

While Brewster’s observations were based on the refraction (transmission) of light through
two optically dissimilar materials, BAM measures the presence of polarized reflected light (Fig 21.) For the measurement of an organic film at an air-water interface where the supporting surface
is water (n2 ≈ 1.33) and the absence of film is air (n1 ≈ 1.00), the angle of incidence is set to
Brewster’s angle (53°), and no reflection occurs. Application of an organic film at the interface
(n ≈ 1.5), changes the ratio between the refractive indices of the air-water interface and reflection
occurs.

Figure 2-1: Description of the application of Brewster's law for the visualization of the
formation dynamics of Langmuir monolayers at an air/water interface.

19

Because the incident radiation is polarized, observed contrast in the image does not necessarily
imply differences in refractive indices between materials. The observed contrast could be due to
changes in the reflected polarization ellipticity induced by the material at the interface as described
by Fresnel.25

BAM Instrumentation. The instrumentation used for the collection of BAM images is
comprised of a diode laser, a microscope objective, and CCD camera (Fig. 2-2). Since the
refractive index for a given medium changes as a function of wavelength, the BAM relies on a 50
mW single wavelength diode laser (658 nm). The angle of incidence can be changed between 52-

Figure 2-2: Schematic of the Brewster Angle Microscope (BAM) system.
57° with a resolution of 0.001° allowing a variety of interfaces to be examined. The polarizer is a
motorized Glan-Thompson prism with a polarizing resolution of 0.001°. The motorized polarizer
ensures an orthoganol plane of incidence relative to the sample surface thereby minimizing
background reflection and increasing observed contrast. The detection optics are comprised of a
10x microscope objective (Olympus, LMPLANFl), analyzer and CCD camera. The microscope
objective allows for a 720x400 Οm field of view with an angular resolution (θ) based on the

20

Rayleigh criterion (eqn. 2-2), where Îť is the wavelength of light (658 nm) and D is the diameter of
the aperture.26-27

θ = 1.220
∆l =0.61

Îť
D

2-2

Îť

NA

By multiplying the radial resolution (θ) by the distance from the object being resolved, the effective
lateral resolution can be described by eqn. 2-2, where NA is the numerical aperture of the objective
(0.25), giving an approximate spatial resolving power of ca. 1.6 Îźm. The motorized analyzer has
a resolution of 0.001° and allows for the correction of induced ellipticity of the initially p-polarized
beam from reflection of the film surface. Finally, the CCD camera provides an image with
1360x1024 px resolution at a frame rate of 20-35 fps.

2.3

Measuring Diffusion: Time-Resolved Fluorescence Spectroscopy
There are a variety of spectroscopic techniques currently used to measure the diffusional

properties of both intact cellular membranes and supported bilayer systems. The most widely
employed methods rely on the use of a fluorescent probe molecule. In this section, information on
the fluorescence microscopy techniques used to examine the diffusion behavior of a chromophore
in a model cell membrane is provided. A brief explanation of the theory behind both the rotational
and translational diffusion measurement techniques is described. The associated instrumental
design and analysis of the observed data is described in detail.

Rotational Diffusion using Fluorescence Anisotropy Decay Imaging (FADI).

The

measurement of steady-state and, specifically, time resolved anisotropy offers the possibility to
study the chemical processes that affect molecular scale reorientation behavior. The process for
21

measuring fluorescence anisotropy relies on photoselective excitation of the chromophores by
polarized light. Photoselective excitation is the preferential excitation of chromophores with
transition moments (electronic dipole) aligned parallel to the electric vectors of the exciting
photons. The transition moment of a chromophore occurs at a defined orientation with respect to
the molecular axis coordinate system. In a fully isotropic solution the transition moments of the
chromophores are oriented randomly. Upon excitation, only the chromophores with absorption
transition moments in the same plane as the polarization of the incoming light are excited.
Emission of light also occurs at a polarization that is fixed relative to the molecular coordinate
system. By measuring the intensity of fluorescence transients both parallel I||(t) and perpendicular
I⊥(t) to the initial excitation polarization over time, the anisotropy of the chromophore can be
measured (eqn. 2-3). The relative angle between the absorption transition moment and emission
moment determines the maximum anisotropy (r0).
R (t ) =

I  (t ) − I ⊥ (t )

2-3

I  (t ) + 2 I ⊥ (t )

There are a variety of physical phenomena that can contribute to an observed anisotropy
decay. The most common causes for an observed anisotropy decay are due to rotational diffusion
and resonant energy transfer.28-34 Because of the sensitivity of the FADI instrument, and the low
concentration of chromophore used, the anisotropy decay signal is dominated by the rotational
diffusion of the fluorescent probe molecule. Measuring the change in anisotropy from the time of
the absorption event to the time of the chromophore emission event provides information on the
average relative angular displacement of the fluorescent probes. In typical fluids, over the
timespan of 50-100 ps, most chromophore ensembles have exhibited rotational motion and the
population of the excited chromophores is closer to being randomized relative to the time of initial
excitation. The anisotropy decay time (𝜏𝜏OR) is inversely related to the diffusion constant (D).
22

Randomization due to rotational diffusion by Brownian motion, is dependent on the temperature
(T), viscosity (Ρ), and volume (V) of the rotating molecule and described by the Debye-StokesEinstein (DSE) equation (eqn. 2-4).35-40 The model was originally derived for spherical molecules
in a homogenous continuum solvent system. The addition of an adjustment for the shape factor of
a nonspherical molecule (S), and the consideration of frictional interactions between solvent and
solute (f) brings the DSE model into good agreement with experimental observations for a
reasonably broad range of experimental systems.41-43

τ=
OR

ΡVf
1
=
6 D k BTS

2-4

The functional form of the anisotropy decay is dependent upon the specific type of
fluorophore under investigation. Below, we describe expected functional forms for an ellipsoidal
rotor free to rotate about all axes, then discuss the case of a tethered rotor with rotation limited
about two axes.
For a free rotor in solution, the anisotropy decay function R(t) can contain up to five
exponential decays, but for most systems, 1-2 decays are observed.29 By referencing a molecules’
Cartesian coordinate system, Chuang and Eisenthal have related R(t) to the Cartesian components
of the rotational diffusion constant (Dx,Dy,Dz).29 The rotational diffusion constants are based on
the rotation of an ellipse where rotation about the major axis (prolate spheroid) is defined by
Dx>Dy=Dz and rotation about the minor axis (oblate spheroid) is defined by Dz>Dx=Dy. By
defining the conjugated fluorophore π-system as the x-y plane, and the absorption transition
moment as the x-axis, a transition from the ground to first excited singlet state (S0 → S1) yields
excitation and emission transition moments nominally parallel to one another. By modeling the
molecule as an ellipsoid and defining the Cartesian coordinates in the previously described
manner, the anisotropy decay function can be described by eqns. 2-5 and 2-6.29
23

=
R ( t ) 0.1e(

−( 2 Dx + 4 Dz )t )

R ( t ) = 0.4e( −6 Dz t )

+ 0.3e(

−6 Dz t )

(oblate)

2-5

(prolate)

2-6

For cases when diffusional motion of the rotor is limited about less than the three
independent axis, the rotor is said to be hindered, and the hindered rotor model applies. The model
is most commonly used for systems in which the fluorescent probe is chemically bound to either
the headgroup or alkyl chain of the lipid molecule. The anisotropy decay function for the hindered
rotor model is described by eqn. 2-7.28, 44

R(t ) = R(∞) + ( R ( 0 ) − R(∞))exp (−t

τ HR )

2-7

In this formalism, the rotor movement is constrained to move within a cone of confinement with a
semi-angle defined as θ0. The rotation of the chromophore about the tethering bond within the
cone gives the probe a characteristic wobbling diffusion constant (DW). The relationship between
the anisotropy decay time (𝜏𝜏HR), the wobbling diffusion constant and cone angle is given by eqn.
2-8.

τ HR =

7θ 02
24 DW

2-8

The cone angle (θ0) can be determined from the infinite-time anisotropy R(∞) and zero-time
anisotropy R(0) according to eqn. 2-9.44
R (∞ )
=
R (0)

1
P2=
( cosθ )  cosθ0 (1 + cosθ0 )
2


2

2

2-9

Fluorescence depolarization is an interaction that is described by the second-order Legendre
polynomial. For any condition where the angle θ0 is zero, the interaction vanishes eqn. 2-10.45
P2 ( cosθ ) = 0

2-10

24

The so called “magic angle” corresponding to this relationship for linear spectroscopy is therefore
defined as 54.7°. For fluorescence depolarization measurements, observation of a cone angle with
a value of the magic angle, therefore corresponds to complete randomization of the chromophore
about the tethered bond.

FADI Instrumentation. The instrument used to acquire fluorescence anisotropy decay
images (FADI) is based on the combination of a time correlated single photon counting (TCSPC)
laser system coupled to an inverted confocal laser scanning microscope (CLSM) (Nikon Eclipse
Ti-U), shown in Fig. 2-3. The light source for this instrument is a synchronously pumped cavity
dumped dye laser (Coherent 702) excited by the output of a passively mode locked Nd:YVO4 laser
(Spectra Physics Vanguard). The laser produces both 355 nm and 532 nm light with 13 ps pulses
at a repetition rate of 80 MHz and 2.5 W average power. The mode locked laser pumps a pair of
dye lasers synchronously which can be tuned from 430 nm to 850 nm. The dye lasers are cavity
dumped to control the repetition rate. The output of the dye laser is characterized by a repetition
rate of 80 kHz – 80 MHz and ~ 5 ps pulses at an average power at the sample of less than 0.5 mW.
For the work presented in this dissertation, the dye laser output is tuned to the excitation maximum
of the chromophore in use. The pulsed excitation light is then passed through a polarizer selected
to an angle of 0 degrees, next through a set of high-pass filters, and finally into the CLSM.

25

Figure 2-3: Schematic for the Fluorescence Anisotropy Decay Imaging (FADI)
instrument.
The CLSM combines high resolution optical imaging with depth selectivity. The CSLM
is based on a typical optical microscope, which uses a laser source instead of the often used
broadband lamp. The laser is focused on the sample, and the image is acquired pixel-by-pixel by
collecting the emitted photons. Detection of time-resolved data is achieved with a polarized dual
channel confocal scanning instrument (Becker & Hickl DCS-120) attached to an output port of the
microscope and controlled by a galvo-drive unit (Becker & Hickl GDA-120). The confocal
scanner is equipped with a polarizing beam splitter and two avalanche photodiode detectors (APD)
(ID-Quantique ID100) for the acquisition of fluorescence lifetime and anisotropy decay images.

26

Polarized fluorescence transients are recorded using time-correlated single photon counting
(TCSPC) detection electronics (Becker & Hickl SPC-152, PHD-400N reference diode).
The TCSPC detection electronics are comprised of two constant fraction discriminators
(CFD), and electrical delay (DEL), Time-to-Amplitude Converters (TAC), an amplifier between
the TAC and the Analogue to digital converter (ADC), and finally to the system memory. The
principle of TCSPC is the detection of the arrival time difference between the emission photons
from the sample with respect to the reference signal from the source (Fig 2-4).

Figure 2-4: Visual representation of the photon counting scheme for time correlated single
photon counting (TCSPC)
The system electronics can be compared to a stopwatch, where the time difference between
the START signal (from the emission photon) and STOP signal (from the excitation pulse) is
measured as an increase in the memory value of a count vs. time difference histogram. For
anisotropy measurements, both parallel and perpendicular transients are measured simultaneously
on separate sets of TCSPC electronics. The combination of the pulsed dye laser source and TCSPC
electronics gives this system an instrument response function of less than 100 ps FWHM.

27

Translational Diffusion by Fluorescence Recovery After Photobleaching (FRAP).
Fluorescence recovery after photobleaching (FRAP) was introduced initially as a means to
measure the translational behavior of fluorescent probe molecules in cellular membranes.46 In a
typical FRAP experiment, the fluorescence of a probe molecule is initially measured over a period
of time (seconds) to obtain a baseline fluorescence intensity. A brief, high intensity, light source
is used to photo-bleach a confocal region of interest containing a subset of the fluorescently labeled
probe molecules. The laser is quickly attenuated and used to measure the fluorescence recovery
as the unbleached molecules diffuse into the region of interest, and the photobleached molecules
diffuse out. The recovery of the fluorescence intensity is strongly dependent upon the translational
behavior of the mobile probe molecules. For a system where the probe molecule is free to diffuse
and does not undergo binding events, the probe is in a purely diffusion limited regime. We first
describe the fitting of the FRAP response curve to a system absent of any binding interactions
(pure diffusion), then describe the fitting and extrapolation of both diffusion and binding
information from a single FRAP response curve.
A recovery curve for a free chromophore (perylene) in a supported lipid (DMPC) bilayer
on a phosphorylated glass surface is shown in figure 2-5. Recovery curves are typically normalized
by the Phair double normalization method to account for acquisition bleaching (eqn. 2-11).47
=
I norm (t )

I ref − pre
I ref − I background

⋅

I frap (t ) − I background

2-11

I frap − pre

For images gathered on supported lipid membranes, the field of view consists of
fluorescence from the chromophores of interest and the background signal cannot be measured
(IBackground). The reference is determined as the region over the entire field of view (Iref). The
FRAP signal from the region of interest (Ifrap) are normalized to both the initial (pre-bleach) signals
of the region of interest (Ifrap-pre) and reference (Iref-pre) to force values between 0 and 1.
28

Figure 2-5: Reference, signal, and normalized response FRAP curve of perylene in a
solid supported DMPC bilayer.
While acquisition bleaching is often unavoidable, the contribution can be minimized by
keeping the attenuated laser intensity low. The time to recover to half of the maximum is termed
the characteristic “diffusion time” (τD) and is directly proportional to the area during the bleaching
event. If the process is diffusion limited, the diffusion time can be related to the translational
diffusion constant (DT) and bleaching radius (ω) by eqn. 2-12.

τD =

ω2

2-12

4 DT

The functional form of the FRAP curve in the diffusion limited regime is described by the
Soumpasis function (eqn. 2-13).48

=
I (t ) e

 −2τ D 
 t 



  −2τ D 
 −2τ D
 I 0  t  + I1  t


 





2-13

29

where I0 and I1 are the modified Bessel functions of the first kind to the zeroth and first order,
respectively.48 Changes to the functional form for a variety of diffusion constants and bleaching
radii (spot sizes) in comparison to the fit raw data (figure 2-5) are shown in figure 2-6. The
convenience of this function is that the translational diffusion constant can be determined by fitting
a single parameter (τD).

Figure 2-6: Functional form for FRAP curves based on the Soumpasis model for various
diffusion times “τD” (s) (left) and spot sizes (μm) (right). The best fit for a recovery curve
of perylene in a solid supported DMPC bilayer is shown with the residuals below.
For probe molecules that undergo interactions with the surrounding medium or supporting
surface, care must be taken to properly fit the acquired data to the proper model. We begin by
describing the equations used to analytically solve the diffusion constant for a single binding
reaction. A general model for a single binding reaction is given by eqn. 2-14.
kon
F +MB

2-14

koff

30

where F =[f] represents the concentration of free diffusing molecule, M = [m] represents the
concentration of an available binding site, such as a phosphonate on the supporting surface, and B
= [b] represents the concentration of bound complex between the chromophore and the binding
site. Assuming the system has reached equilibrium prior to the bleaching event, and that the
binding sites are immobile, the recovery can be described by a set of coupled reaction-diffusion
equations49 eqn. 2-15:

∂f
*
= Dt ∇2 f − kon
f + koff b
∂t
∂b
*
= kon
f + koff b
∂t

2-15

where Dt is the diffusion constant of the chromophore, k*on = k*on[m] is the pseudo first-order rate
constant derived by assuming that the M = [m] is constant, and ∇2 is the Laplacian operator. For
the assumptions to hold, it is important to allow time for the supported film to achieve equilibrium
both prior to, and during, imaging. Confirmation of an equilibrium state can be achieved by
measuring changes in the fluorescence intensity prior to the bleaching event. Given a system at
equilibrium at the start of the FRAP experiment both F and B are at equilibrium:

df db
= = 0
dt dt
*
kon
Feq = koff Beq

2-16

Assuming normalization of the FRAP curves between 0 and 1, the final recovery of the free and
bound species is equal to:

Feq + Beq =
1

Feq
=

*
koff
kon
B
=
,
eq
*
*
kon
kon
+ koff
+ koff

2-17

For a system in a reaction-dominant regime, where diffusion is very fast compared to
binding and the timescale of the FRAP measurement (kon*w2/DT << 1) the recovery can be fit to a
31

simple exponential form. Bulinski et al. initially solved a general solution relating the dissociation
rate constant (koff) to the functional form of the FRAP curve and Sprague et al. have further related
the recovery to both koff and kon* (eqn. 2-18).49-50
I norm ( t ) = 1 − Beq e

− koff t

2-18

One important consideration concerning this function is the lack of dependence on the
bleaching spot size. Given the change of the functional form for a variety of spot sizes with the
same diffusion constant, additional experiments must be done to determine whether the system is
in a fast diffusion vs. binding regime. The dissociation rate constant (koff) is inversely related to
the square of the bleaching radius. Thus, by plotting the determined koff rate vs. a variety of spot
sizes a linear response is expected. A nonlinear response suggests that the system is not in a
reaction-dominant regime and will be better fit and described by the full reaction-diffusion model.
Sprague et al. have shown one way to analytically solve the reaction-diffusion differential
equations. The inverse Laplace transform of the differential diffusion equations can be performed
using the matlab function invlap.m on the analytical equation:
*
 1 Feq

kon
Inorm (=
t)   −
⋅ 1 − 2K 1 ( qω ) I1 ( qω )  ⋅  1 +

p + koff

 p p
−1

  Beq
 − 
  p + koff

 
 
 

2-19

where Inorm(t) is recovered by taking the inverse Laplace transform, Feq and Beq are related to k*on
and koff by eqn. 2-17, K1 and I1 are the modified bessel functions, ω is the radius of the bleaching
spot and q is described by eqn. 2-20.51
q
=

 p

 Df

*

kon
1+


p + koff






2-20

While eqn. 2-20 provides solutions for k*on and koff and the translational diffusion constant
(Df), initial parameters for the fit are underdetermined. To estimate the initial parameters, the

32

recovery curve is fit to both the Soumpasis function (eqn.2-13) and the reaction dominant model
(eqn.2-18). The full reaction-diffusion model is best used as a model for a tethered chromophore
where the assumptions of the reaction-dominant and Soumpasis models do not hold.

FRAP Instrumentation. The instrument used to perform FRAP measurements is based on
an inverted confocal laser scanning microscope (CLSM) (Nikon C2+ system) equipped with a four
diode laser bank (Nikon LU-N4, 405 nm, 488 nm, 561 nm, 640 nm). The microscope is equipped
with a LED illuminator for the collection of steady state images, a motorized sample stage, and a
3 PMT detector (400-700 nm) (Fig. 2-7).

Figure 2-7: Schematic for the Fluorescence Recovery After Photobleaching (FRAP)
instrument.

33

Data obtained from the combination of BAM, FRAP, and FADI instruments are used in
subsequent chapters of this dissertation to understand how the fluidity, adhesion, and heterogeneity
of supported lipid membranes are influenced by a variety of exogenous factors.

34

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35

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Perrin, F., Brownian movement of an ellipsoid (II). - Free rotation and depolarisation of
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Measurement by Analysis of Fluorescence Photobleaching Recovery Kinetics. Biophys J
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Phair, R. D.; Gorski, S. A.; Misteli, T., Measurement of dynamic protein binding to
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Soumpasis, D. M., Theoretical-Analysis of Fluorescence Photobleaching Recovery
Experiments. Biophys J 1983, 41 (1), 95-97.

49.

Sprague, B. L.; Pego, R. L.; Stavreva, D. A.; McNally, J. G., Analysis of binding reactions
by fluorescence recovery after photobleaching. Biophys J 2004, 86 (6), 3473-3495.

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Bulinski, J. C.; Odde, D. J.; Howell, B. J.; Salmon, T. D.; Waterman-Storer, C. M., Rapid
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Hollenbeck, K. J. INVLAP.M: A matlab function for numerical inversion of Laplace
transforms by the de Hoog algorithm, 1998.

39

CHAPTER 3: LIPID ADLAYER ORGANIZATION MEDIATED
BY A LIQUID OVERLAYER

Adapted with permission from:
S. M. Baumler and G. J. Blanchard; Lipid adlayer organization mediated by a liquid overlayer.
Spectrochimica Acta Part A: Molecular and Biomolecular Spectroscopy 98 (2012) 429–435.

40

3.1

Abstract
Formation of a chemically bound 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC)

monolayer on modified gold and silica surfaces, and changes in the organization of the interfacial
lipid layer associated with immersion in aqueous solution were examined. The interface was
studied using steady state and time resolved fluorescence spectroscopy, water contact angle,
optical ellipsometry measurements, and electrochemical methods. The DMPC adlayer in contact
with air forms a relatively well organized interface that mediates the rotational motion of perylene.
In the presence of an aqueous overlayer, perylene reorientation becomes more rapid, consistent
with a reduction in the organization of the interfacial lipid adlayer. One implication of this finding
is that the interfacial adlayer is less than a uniform monolayer, which is confirmed by
electrochemical impedance data. These findings underscore the importance of water in mediating
the organization of interfacial lipid adlayers.

3.2

Introduction
The development of chemical sensing strategies has relied heavily on the ability to modify

interfaces to impart chemical selectivity. Among the most important aspects of chemical surface
modification has been the effort to create biomimetic interfaces. The creation of such interfaces
is critical to the development of selective sensors that employ biomolecules such as transmembrane
proteins and certain enzymes. For these reasons, there has been much effort invested in the
creation of supported lipid adlayer structures. Lipid membrane composition and organization are
central to the function of transmembrane proteins, with many important cellular activities requiring
a specific lipid membrane micro-domain in order to allow the transmembrane protein to function.

41

Membrane fluidity, which is inextricably tied to membrane composition, also plays an important
role in allowing processes such as cell adhesion, ion channeling, and cell signaling to operate.1-3
For a biomimetic interface to be useful for sensing or other applications, the interface must
be supported on a solid substrate for chemical signal transduction by electrochemical,
spectroscopic or gravimetric means, and for other applications the presence of a solid substrate is
required to impart the requisite physical robustness to the interface. There have been a number of
strategies put forth for attaching biomimetic interfaces to surfaces.4-6 Simple physisorption
processes such as those responsible for the formation of Langmuir-Blodgett films on solid supports
are typically not robust enough to provide the properties needed for most biomimetic interfaces.
Several strategies offer abundant chemical and physical integrity, but the issue of adlayer fluidity
remains to be addressed fully for such systems.7-14
Extensive attention has been paid to the preparation of air stable supported membrane
interfaces because of the fragility of typical bilayer systems and the potential utility of biomimetic
systems able to withstand direct exposure to the ambient environment.15-16 Preparing a robust
system would have use in bio-electronics and would allow for a stable model to investigate
functional proteins. Because of the requirement of a robust system and the intended applications
for such a system, three common attributes are sought when preparing bound interfacial systems.
The systems should be physically robust, easily and reproducibly prepared, and be sufficiently
uniform to allow subsequent processing for specific applications.
When lipid adlayers are deposited onto, or bound to an interface in a layer-by-layer manner,
there is at least one step in the process that necessarily presents a non-polar interface for subsequent
processing. It is this step in the formation of a biomimetic interface that we are concerned with in
this work. Specifically, the creation of a non-polar interface may proceed with substantial order.

42

Due to the processing of such interfaces in the laboratory environment, it is often impossible to
prevent the exposure of the interface to adventitious water, and sometimes aqueous solutions are
used in the deposition of subsequent layers. The interaction(s) between the nonpolar region of the
interfacial layer and water could, in principle, alter the organization of the nonpolar region of the
layer prior to or during subsequent adlayer deposition steps. We have formed a phospholipid
adlayer on polar surfaces to produce an interface with its non-polar region being exposed to the
environment. Examination of the non-polar region of the interface as a function of its exposure to
the ambient environment and aqueous solution demonstrates that the organization of the interface
is indeed sensitive to the presence of bulk water. Ionically bound phospholipids are capable of
exhibiting significant organization and have been shown to withstand ambient environmental
conditions without loss of structural integrity.14, 17-19 The presence of bulk water, however, has a
substantial effect on the organization of the adlayer. The motional dynamics of a non-polar
chromophore sequestered within the lipid adlayer reveal a change in acyl chain organization that
gives rise to increased motional freedom when the interface is exposed to water.

3.3

Experiment
Materials. Acetonitrile, chloroform, ethanol (100%), toluene, TrisÂŽ buffer, 2,4,6-collidine,

(3-aminopropyl)

dimethylethoxysilane

(APDMES),

6-mercapto-1-hexanol,

phosphorus

oxychloride, potassium ferrocyanide trihydrate, potassium chloride, perylene, and zirconyl
chloride octahydrate were obtained from Sigma Aldrich in the highest purity grade available and
were used as received. The phospholipid 1,2-dimyristoyl-sn-phosphatidylcholine (14:0 DMPC)
was purchased from Avanti Polar Lipids, Inc., and used without further purification. Water (18
MΊ) was obtained from a Milli-Q Plus water purification system and used for all experiments.

43

Figure 3-1: Chemical Structures of 1,2-dimyristoyl-sn-phosphatidylcholine (DMPC) and
Perylene.

Metal-Phosphate Monolayer Synthesis. Silica and gold substrates were prepared by a
procedure described elsewhere.20-22

Silica substrates were immersed in piranha solution

(1H2O2:3H2SO4, Caution! Strong oxidizer!) for 20 min., rinsed with distilled water, then ethanol
and dried under a N2(g) stream. The silica substrates were then immersed in a 0.5% v/v solution
of (3-aminopropyl)dimethylethoxysilane in dry toluene for ca. 12 hrs. Gold substrates (ca. 200
nm Au vapor-deposited on Si(100) with a 20 nm Ti adhesion layer) were treated with 10 mM 6mercapto-1-hexanol in ethanol for ca. 12 hrs. Following the reactions, both substrates were rinsed
with ethanol and dried under a N2(g) stream. The APDMES adlayers on silica and gold substrates
were then exposed to POCl3 (0.4 mL) in dry acetonitrile (10 mL), catalyzed with 2,4,6-collidine
(0.4 mL) for 3 hrs. at 20°C. The gold substrates were then rinsed with ethanol and distilled water
and the silica substrates were rinsed with acetonitrile. Both substrates were dried with a N2(g)
stream. Zirconation of the phosphate-terminated silica and gold substrates was achieved by
immersion in 5 mM ZrOCl2 in 60:40 ethanol/water solution for 12 hrs. The resulting zirconated

44

substrates were rinsed with water and dried under a N2(g) stream, and reacted with a solution
containing unilamellar phospholipid vesicles.

Figure 3-2: Steady state excitation (S1 ← S0, solid line) and emission (S0 ← S1, dashed
line) of perylene in DMPC vesicles prior to formation of the lipid monolayer.

Vesicle Preparation. For each substrate, 10 mg of the phospholipid 14:0 DMPC was mixed
with 50 Îźg of perylene (0.5 wt%) (Fig. 3-1). The chloroform was then evaporated and the resulting
solid mixture dried in vacuo. Tris® buffer (10 mM, pH 7.5) was prepared with 18 MΩ water and
added to the sample to make the final lipid concentration 1 mg/mL and a perylene concentration
of 20 ÂľM. The lipid and chromophore mixture was processed through five freeze-thaw-vortex
cycles to ensure thorough mixing.15 Each cycle consisted of a 5 min. freeze by immersion in N2(l),
5 min. thaw in a 60°C water bath, and a 2 min. vortex. After five such cycles, the samples were
extruded eleven times through a NucleoporeÂŽ polycarbonate membrane (Whatman) with 400 nm

45

diameter pores using a mini-extruder apparatus (Avanti Polar Lipids, Inc.) to produce unilamellar
vesicles.16-17 Figure 3-2 contains the normalized steady state excitation and emission spectra of
perylene contained in the extruded vesicles. Previous work has shown that the steady state spectra
of perylene are insensitive to the phase or structural format (i.e. vesicle, supported bilayer) of the
lipid bilayer medium.23-24

Time resolved spectroscopic data provide more environmental

information for this chromophore.

Lipid Adlayer Formation. Phospholipid adlayers were formed by spontaneous fusion of
unilamellar lipid vesicles onto the zirconated substrate surfaces (Fig. 3-3).25 A TeflonÂŽ flow cell
constructed in-house was used to ensure consistent contact between the substrate and vesicle
solution during the phospholipid adlayer formation process.17 The substrate was conditioned with
a TrisÂŽ buffer solution introduced at a rate of ca. 5 mL/min. The vesicle-containing solution was
introduced at the same rate until the buffer was displaced by ca. 4 mL of solution. The flow cell
inlet and outlet were then sealed and the system was allowed to react for 2 hrs. Water (18 MΩ)
was subsequently flowed through the cell to rinse the substrates, which were then removed from
the flow cell and mounted vertically in a loosely capped vial to dry.

46

Figure 3-3: Idealized diagram of the prepared supported lipid layer. The films are
composed of the (a.) gold or silicon support, (b.) phosphorylated APDMES (c.)
zirconation layer, and (d.) DMPC layer.

Steady State Fluorescence Measurements. Excitation and emission spectra were recorded
for our samples (Jobin Yvon Fluorolog-3) to verify the presence and band positions of the
incorporated perylene chromophore. Spectral resolution was set to 3 nm for both the excitation
and emission monochromators.

Time Resolved Fluorescence Measurements.

Fluorescence lifetime and anisotropy

measurements were performed using an instrument that has been described in detail in Chapter 2
and we recap its salient features below. The light source for the system is a CW passively mode-

47

locked Nd:YVO4 laser (Spectra Physics Vanguard) that produces 13 ps pulses at 1064 nm at a
repetition rate of 80 MHz. The third harmonic output of this laser excites a cavity-dumped dye
laser (Coherent 702-2). The output of the dye laser is typically 5 ps fwhm pulses at a repetition
rate of 4 MHz, determined by the cavity dumping electronics (Gooch and Housego). Stilbene 420
dye (Exciton) used to excite the samples at 440 nm. The pulses from the dye laser are divided,
with approximately one half going to a reference diode (Becker and Hickl PHD-400-N), and the
other half going to the sample. The sample is excited with the vertically polarized light pulses and
emission is collected at polarizations parallel and perpendicular to the excitation polarization. The
polarized emission data are collected simultaneously using subtractive double monochromators
(Spectral Products CM-112) equipped with micro-channel plate PMT detectors (Hamamatsu
R3809U-50) detectors.

Detection electronics resolve the parallel and perpendicular signals

separately (Becker and Hickl SPC-132). Collection wavelength, data acquisition, and detector
bias are all controlled using a program written in-house (National Instruments LabVIEWÂŽ).

Electrochemical measurements.

Data were acquired using a CH Instruments 650

electrochemical bench. The solution used to characterize the interfaces was 0.1 M KCl. Cyclic
voltammograms (CV) were recorded with 3 independent scans cycled two times each at a scan
rate of 0.1 V/s. The substrates were scanned from -0.1 V to 0.4 V vs. Ag/AgCl using a Pt counter
electrode. Impedance measurements were collected using the same probe, and counter and
reference electrodes. A frequency range of 0.1 Hz to 105 Hz was used at an AC amplitude of 5
mV. The center voltage was set as E0, determined by the individual CV scans (0.26 V vs
Ag/AgCl).

48

3.4

Results and Discussion
The design and demonstration of physically and chemically robust biomimetic interfaces

has generated substantial research interest for both fundamental and applied reasons. One aspect
of this work has been the chemical binding of interfacial adlayers, with an ultimate goal of utilizing
such systems for chemical sensing applications. Because of the amphiphilic nature of lipids, the
details of how a supported lipid bilayer is formed can have a substantial impact on the organization
and properties of the interface. Specifically, for systems formed one lipid layer at a time with the
acyl chains exposed, the lipid layer is at some point exposed to a polar (aqueous) liquid overlayer.
We are interested in this work in determining the consequences of this exposure on the
organization of that layer. We consider the thickness, uniformity and permeability of an interfacial
lipid monolayer as well as its intralayer organization.

Interfacial thickness and hydrophobicity.

Optical ellipsometry and contact angle

measurements were performed to confirm phospholipid adlayer formation and to determine the
monolayer or bilayer status of the substrate. Adlayers formed on the gold substrates were
determined to have a thickness of 48 ± 10 Å, and water contact angles of 84° ± 9. The silica
substrates exhibit the same contact angle (95° ¹ 5) to within the uncertainty of the measurements.
The ellipsometric and contact angle data are consistent with the formation of a DMPC monolayer
on both support materials.17 We note that the ellipsometric thickness appears to be somewhat
thicker than a hydrated DMPC layer, but the value of 48 ± 10 Å also contains the thickness of the
phosphate and zirconated APDMES adlayer (ca. 15 Å). The data that we report for each technique
represent an average of at least three determinations per substrate. Each substrate was measured
at least three times at different locations, and the results were averaged. The contact angle

49

measurements follow the same procedure and were measured after allowing the droplet to ‘settle’
for ca. 30 sec. before the measurement was taken.

Chromophore motion within the lipid layer.

We have measured the fluorescence

anisotropy decay of perylene imbedded in the bound DMPC adlayer when it is exposed to air and
immersed in water. Anisotropy decay measurements contain information on the motional freedom
of the chromophore. The orientational anisotropy decay function, R(t) eqn. 3-1, is the normalized
difference between polarized emission transients I||(t) and I⊥(t), where the polarizations are relative
to the vertically polarized excitation pulse.
R (t ) =

I  (t ) − I ⊥ (t )

3-1

I  (t ) + 2 I ⊥ (t )

The functional form of R(t) is related to the chemical information contained in these data. Chuang
and Eisenthal have derived the relationship between the polarized emission transients, the
orientations of the chromophore absorbing and emitting transition moments, and the Cartesian
components of the rotational diffusion coefficient, D eqn. 3-2.27

D=

1
3

(D

x

+ Dy + Dz )

3-2

In their model, the volume swept out by the reorienting molecule is described as a prolate
or oblate ellipsoidal shape, with control over the aspect ratio of the ellipsoid of rotation. While
the Chuang and Eisenthal equations provide for up to five exponential decay components in R(t),
either one or two decay components are observed experimentally. Depending on the number of
decay components and the relative orientations of the excited and emitting transition moments,
prolate and oblate ellipsoidal rotor shapes give rise to different functional forms of R(t). The S1
← S0 transition dipole moment of perylene lies in the long in-plane axis of the chromophore, which

50

we designate the x-axis. For perylene the y-axis is the short in-plane axis, and the z-axis is
perpendicular to the π-system plane. For rotation as a prolate rotor with x-axis rotation being
dominant (Dx > Dy = Dz), R(t) is given by eqn. 3-3,
=
R(t ) 0.4 exp(−6 Dz t )

3-3

For rotation as an oblate (z-axis dominant) rotor (Dz > Dx = Dy), a two-component exponential
decay of R(t) is expected (eqn. 3-4),
R(t =
) 0.1exp(−(2 Dx + 4 Dz )t ) + 0.3exp(−6 Dx t )

3-4

Figure 3-4: Induced orientational anisotropy decay data for perylene in DMPC
monolayer on SiOx substrates, (a.) exposed to air and (b.) exposed to an aqueous
overlayer. The solid lines are the best fit, single exponential decay function. Insets
represent residuals of fit.
We observe a single exponential anisotropy decay for the measurements we report here, consistent
with perylene reorienting as a prolate rotor (eqn. 3-3, Fig. 3-4). It is useful to place this result in
context. Perylene is capable of reorienting either as a prolate rotor or as an oblate rotor, depending
on its local environment.24, 28 In n-alkanes, perylene reorients as a prolate rotor28 and in other

51

environments, such as within a lipid bilayer, perylene has been shown to reorient as an oblate
rotor.24 The fact that we observe a single exponential decay in this work underscores the difference
between the characteristic organization within a lipid monolayer and that within a lipid bilayer
structure.
The data presented in Table 3-1 reveal a statistically discernible difference between the
reorientation dynamics of perylene in lipid monolayers that are in contact with air and those
immersed in water. Specifically, the reorientation time constant of perylene in bound lipid
adlayers in contact with air is ca. 40% slower than when the monolayer is immersed in water. This
finding indicates that the lipid monolayer in contact with air is better able to form organized
structure(s) within its nonpolar region. We assert that bulk water in contact with the nonpolar acyl
chains of the lipid adlayer disrupts the organization of the lipid acyl chain region. It is important
to recognize that, regardless of whether we have a full or partial lipid adlayer, the perylene
chromophore will reside in the most non-polar portion of the interface, which is the acyl chain
region of the lipid adlayer. Our rotational diffusion data thus indicate that exposure of the interface
to water decreases the order that exists within the phospholipid acyl chain region(s) of the interface.
We will consider the implications of this finding after discussing the chromophore reorientation
data in detail.
Table 3-1: Reorientation time (𝜏𝜏OR), Rotational Diffusion Constant (D), and Calculated
Local Viscosity (Ρ) determined by Local Environment for DMPC Bound to a Modified
Silica Substrate.
Ambient Adlayer
Aqueous Adlayer
Substrate
D (MHz)
Ρ (cP)
τOR (ps) D (MHz) η (cP)
τOR (ps)
S1
351 Âą 72 487 Âą 90 4.5 Âą 0.9
101 Âą 6
1651 Âą 103 1.3 Âą 0.1
S2
281 Âą 11 594 Âą 22 3.6 Âą 0.1
183 Âą 34
932 Âą 156 2.3 Âą 0.4
S3
288 Âą 77 604 Âą 132 3.7 Âą 1.0
105 Âą 32
1733 Âą 632 1.3 Âą 0.4
Average
307 Âą 39 562 Âą 82 3.9 Âą 0.5
130 Âą 46
1439 Âą 297 1.7 Âą 0.6
Errors are ¹ 1σ for at least 3 individual determinations, on each substrate, and in each
environment.

52

In addition to the shape of the volume swept out by the rotating chromophore, it is also
useful to relate the experimental data to physico-chemical properties of the local environment. The
modified Debye-Stokes-Einstein (DSE) equation (eqn. 3-5) provides a means of relating the
rotational diffusion constant, D (eqn. 3-2), to the local viscosity of the medium.29-32
D=

k BTS
6ΡVf

3-5

This model is well established for a variety of solution phase systems, where Ρ is the
viscosity of the medium surrounding the rotating chromophore, V is the chromophore
hydrodynamic volume (225 Å3),33 S is a shape factor relating to the ellipsoidal shape of the
chromophore (S = 0.69)31-32 and f is a term to account for frictional contributions to the interaction
between the chromophore and its surroundings (0 ≤ f ≤ 1).30 This model was derived for
reorientation of dipolar species in a continuum medium, and a direct comparison between a
chemically-bound DMPC adlayer on a planar substrate and a bulk liquid solvent requires some
caution. It is nonetheless useful to examine the bound lipid layer using the DSE model because of
the fluid nature of the adlayer acyl chain region. The experimental data demonstrate that perylene
is able to execute rotational motion within the acyl chain environment, but comparison of the
“viscosity” of the adlayer to that of a bulk liquid is tenuous because of the quasi two-dimensional
nature of the adlayer and the fact that it is bound to the interface. Given this caveat, the rotational
motion of perylene in the adlayer acyl chain region exhibits an apparent viscosity that is similar to
that seen for alkanes that are similar in length to the acyl chains (Table 3-1). The viscosity of
tridecane, tetradecane, and hexadecane are 1.7 cP, 2.1 cP, and 3.0 cP, respectively, at 25°C.34
What is most significant about these data is, however, that the immersion of the surfacebound adlayer into water gives rise to a substantial decrease in the perylene reorientation time
constant. This substantial change in the rotational diffusion constant of the chromophore is

53

consistent with a loss of organization in the acyl chain region of the adlayer. It is unfortunately
not possible to elucidate possible changes in the aspect ratio of the ellipsoid of rotation that
describes perylene reorientation in this adlayer, but such a determination is not possible when only
a single exponential anisotropy decay is recovered (eqn. 3-3). The fact that the chromophore
experiences more motional freedom within the adlayer when it is immersed it water invites
speculation on the cause of this effect. Electrochemical data can provide some insight into possible
explanations for this finding.

Interfacial thickness and heterogeneity characterization. Determining the thickness and
extent of surface coverage of a surface-bound adlayer can be challenging. We have measured the
thickness of our APDMES+ZP+DMPC adlayers on Au ellipsometrically to be 48 ± 10 Å, but this
thickness value does not address heterogeneity of the adlayer. We use electrochemical methods
for this purpose. We recognize that the process of vesicle fusion is complex and system-specific,
and in general, that it is not possible to compare lipid adlayers formed on silica and Au surfaces
directly.4-6 The work presented here, however, is somewhat different than most other studies of
surface-bound lipid adlayers. In this work we functionalize both the silica and Au surfaces to form
terminal phosphate moieties, followed by treating those surfaces with Zr4+ prior to exposure to the
phospholipid. While there will certainly be differences, in detail, between the silica and Au
surfaces, the fact that both are terminated with the same chemical functionality provides a greater
correspondence between these surfaces than would be found if no surface modification had been
applied.

54

Figure 3-5: Cyclic voltammograms of a blank Au surface (─), a C12 thiol monolayer (---)
and a DMPC adlayer bound to Au (⋯⋯). All CVs were taken in 0.1M KCl (aq) solution at
a scan rate of 100 mV/s.
The fractional coverage of the surface can be estimated through the capacitance of the
interface. For a uniform adlayer, the capacitance is related to the adlayer thickness through (eqn.
3-6),
C=

Îľ 0Îľ A


3-6

where Îľ is the dielectric constant of the interface, Îľ0 is the permittivity of free space (Îľ0 = 8.854x1012

F/m), A is the area of the electrode and  is the thickness of the interfacial adlayer. For a

heterogeneous structure, however, such a relationship does not hold directly and the total
capacitance measured by cyclic voltammetry (Fig. 3-5) is modeled as distinct contributions from
the covered and bare regions of the interface. The equivalent circuit for such a heterogeneous

55

surface is two capacitors in parallel, with one being related to the adlayer-coated regions and the
other being related to the bare regions (eqn. 3-7),35
Ctot= θ Cml + (1 − θ )Cdl

3-7

where θ is the fractional surface coverage by the adlayer, Cml is the capacitance of the coated
regions and Cdl is the double layer capacitance, characteristic of the bare electrode regions.
Because two quantities in eqn. 3-7 are measurable, Ctot and Cdl, it is not possible using
cyclic voltammetry alone to determine uniquely θ and Cml. For this reason, we have chosen to use
AC impedance measurements to extract information on interfacial coverage and heterogeneity.
AC impedance data is presented in the form of Nyquist plots (Fig. 3-6),36 where the real and
imaginary parts of the measured impedance are plotted against one another as a function of
frequency. For both the DMPC (Fig. 3-6) and the C12SH self-assembled monolayer (SAM, Fig.
3-6), the bare gold substrate (open dots) produces Nyquist plots consistent with a diffusion-limited
reaction. For the C12SH and DMPC adlayers, the Nyquist plots are consistent with partial
monolayer films. The relationship between θ, the fractional surface coverage, and Rct is given by
eqn. 3-8.35
θ = 1−

Rct0
Rct

3-8

where Rct0 is the charge transfer resistance for a bare substrate and Rct is the charge transfer
resistance for the adlayer-containing interface. We obtain a value of Rct0 = 50 Ί from modeling
of the complex impedance data using a Randles equivalent circuit (Fig. 3-6 inset).37-40 Cyclic
voltammetry measurements give values of Cdl = 22 ÂľF for the C12 SAM and 19.5 ÂľF for the DMPC
adlayer, and using these values we obtain Rct = 545 Ί for the C12 SAM and Rct = 105 Ί for the
DMPC adlayer. These data yield θ = 0.91 for the C12SH adlayer and θ = 0.52 for the DMPC

56

adlayer. With the quantity θ, one can determine Cml (eqn. 3-7), which allows estimation of  (eqn.
3-6).

Figure 3-6: AC impedance for a blank Au surface (●), a C12 thiol SAM on Au (■), and a
DMPC adlayer bound to Au (▲). Inset: Equivalent circuit used to extract Rct values
from these data: Ru = uncompensated resistance, Rct = charge transfer resistance, Cdl =
double layer capacitance.
The lower double layer capacitance and lower charge transfer resistance indicate less
complete coverage for the DMPC adlayers than for the C12 SAM. Using eqn. 3-6 to estimate the
thickness, however, leads to values that are too small by a factor of ca. 10. Using eqn. 3-6 and a
dielectric constant of Îľ = 5,41 a value somewhat in excess of what is usually assumed,42 we obtain
thicknesses of 5 Å for the C12SH SAM and 2 Å for the DMPC adlayer. The low electrochemical
estimate for the adlayer thickness is not surprising given that the dielectric response of the adlayer
is an estimate at best and there is some inherent uncertainty in the determination of both C and A.

57

We take the ellipsometric values of thickness to be better approximations of the true coverage of
the surfaces.

Adlayer morphology. The central issue to be considered is the correct physical and
chemical interpretation of the perylene reorientation data, where faster reorientation is observed
for perylene imbedded in DMPC interfaces when those interfaces are immersed in water relative
to the behavior when the interfaces are exposed to air. There are several possible explanations for
these findings. For perylene in the DMPC interface in contact with air, we measure a reorientation
time of τOR = 307±39 ps, consistent with a local viscosity η = 3.9 ± 0.5 cP (Table 3-1). Given that
the reorientation of perylene in a lipid bilayer is consistent with a local viscosity of ca. 10 cP below
the gel to fluid phase transition (24°C) and ca. 7 cP above the phase transition,23 our observation
of ca. 4 cP for perylene in a (sub)monolayer indicates that the DMPC moieties bound to the
interface are relatively close in spatial proximity to one another. This finding is consistent with
the 52% coverage determined electrochemically.
For the same interfacial assembly immersed in water, we recover a perylene reorientation
time constant of τOR = 130 ± 46 ps, corresponding to η = 1.7 ± 0.6 cP (Table 3-1). The fact that
perylene reorients faster in the same interface when it is immersed in water indicates either that
the DMPC-rich regions on the interface are experiencing a significant organizational disruption or
that perylene is being solvated in some manner by water. We believe the latter explanation to be
unlikely given the exceedingly low solubility of perylene in water (~10-8 M). If, as is the case for
the mixing of water and alkanes, a phase boundary exists at the interface of the water and lipid
acyl chains, the question is how the water overlayer alters the lipid acyl chain organization to
increase local free volume. If the thickness of the adlayer is decreased, then the lateral area of the

58

adlayer must be able to compensate for this loss of thickness. It has been shown previously that
the combination of certain substrates and priming chemistry give rise to the formation of islandlike features on the surface of silica substrates.20 Because the DMPC adlayer formation on the
substrates is dependent upon the priming chemistry,17-18 it is likely that the DMPC monolayer is
aggregated, and a surface coverage of 52% would leave space for inter-island regions. The
presence of a water overlayer could, in principle, compress the adlayer, as is suggested by the
ellipsometry data (Table 3-2), and depending on the extent of the compression, such a change
could either increase or decrease the available free volume within the structure.
Table 3-2: Adlayer ellipsometric thickness and electrochemical coverage.
Substrate
Ellipsometric Thickness (Å) Electrochemical Coverage (%)
25 Âą 6
91
C12 SAM
48 Âą 10
52
DMPC/Au
Measurements were performed for two different C12 thiol/Au SAMs and five different
APDMES+ZP+DMPC terminated interfaces on modified gold substrates.
What is not clear from these data is the extent to which the interfacial organization can be
controlled by the composition of the overlayer. There is limited experimental latitude in altering
the identity of the overlayer because of the solubility of perylene in a number of polar and nonpolar
non-aqueous solvents (e.g. methanol, cyclohexane). We have added NaCl (several different
concentrations) to the aqueous solution in which the interface has been immersed and we find no
measurable effect on the reorientation dynamics of perylene. This finding indicates that direct
interactions between the chromophore and liquid overlayer are not responsible for our findings.
Rather, it is the alteration of the DMPC acyl chain organization by the presence of the water
overlayer that mediates the perylene reorientation dynamics. Further, the extent to which the
aqueous phase overlayer influences the organization of the lipid acyl chain region is limited
because the presence of salt in this solution does not lead to organization in the acyl chain region
that is discernibly different than it is for a water overlayer.

59

We are limited in terms of the information that can be extracted from the reorientation data
because we recover single exponential anisotropy decays in all cases. The prolate rotor shape that
produces this decay functionality is related to only one Cartesian component of D and the
measurement is insensitive to changes in the aspect ratio of the ellipsoidal rotor as a function of
local environment. We do consider that the magnitude of the changes we measure in Dz implies
significant disruption is associated with immersion of this interface in water. It is not likely that
the infiltration of even a significant amount of water into the adlayer would give rise to the
observed decrease in effective viscosity. In comparison, the addition of up to 1.6 M ethanol to
aqueous solutions containing 100 nm diameter DMPC vesicles produced only modest changes in
the viscosity of the lipid acyl chain region and essentially no change in the effective rotor shape of
the perylene contained within the bilayer structure.43 Further experimentation will be required to
elucidate in more detail the changes in acyl chain organization that result from exposing bound
lipid monolayers to a water overlayer.

3.5

Conclusions
We have investigated the rotational diffusion dynamics of perylene contained within a

tethered lipid monolayer. The data show relatively restricted chromophore motion in the lipid
adlayer exposed to air and substantially increased motional freedom when the adlayer is immersed
in aqueous solution. The central question is how the water overlayer can interact with the lipid
monolayer to produce a less restrictive local environment for perylene. The dynamics of the
chromophore when the adlayer is immersed in aqueous solution are the same whether sodium
chloride is present in the water or not. The decrease in rotational diffusion time constant of
perylene in the immersed interface is likely associated with a decrease in the extent of organization

60

within the (bound) lipid acyl chains. Such a situation would require a net reduction in local density
of the acyl chains, and this is feasible in light of electrochemical data that indicate ca. 52%
coverage of the lipid monolayer. A more detailed understanding of lipid monolayer morphology
for both air- and water-exposed systems is required. Ellipsometric and contact angle data suggest
a spatially heterogeneous, largely nonpolar interface, but little information is available on the
characteristic length scale(s) of any organized regions on the surface. It is possible that future
studies using more rigid chromophores with different aspect ratios (e.g. DPH)44 may shed light on
the nature of the acyl chain organization in the lipid adlayer, and fluorescence lifetime and
anisotropy imaging data may shed light on the characteristic domain sizes seen for these systems.

61

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30.

Hu, C.-M.; Zwanzig, R., Rotational friction coefficients for spheroids with the slipping
boundary condition. J. Chem. Phys. 1974, 60 (11), 4354-4357.

31.

Perrin, F., Mouvement brownien d'un ellipsoide (I). Dispersion dielectrique pour les
molecules ellipsoidale. J. Phys. Radium 1934, 5, 497.

32.

Perrin, F., Mouvement Brownien d'un ellipsoide (II). Rotation libre et depolarisation des
fluorescences. Translation et diffusion de molecules ellipsoidales. J. Phys. Radium 1936
7,1-11.

33.

Edward, J. T., Molecular volumes and the Stokes-Einstein equation. J. Chem. Ed. 1970, 47
(4), 261-270.

34.

Lide, D. R., CRC Handbook of Chemistry and Physics. CRC Press, Inc.: Boca Raton, FL,
1990-1991; Vol. 71, p 8-39.

35.

Garcia-Raya, D.; Madueno, R.; Sevilla, J. M.; Blazquez, M.; Pineda, T., Electrochemical
characterization of a 1,8-octanedithiol self-assembled monolayer (ODT-SAM) on a
Au(111) single crystal electrode. Electrochim. Acta 2008, 53, 8026-8033.

36.

Pletcher, D., A First Course in Electrode Processes, 2nd Edition. The Royal Society of
Chemistry: Cambridge, UK, 2009.

37.

Randles, J. E. B., Kinetics of Rapid Electrode Reactions. Disc. Faraday Soc. 1947, 1, 1119.

38.

Randles, J. E. B., Kinetics of rapid electrode reactions. Part 2. - Rate constants and
activation energies of electrode reactions. Trans. Faraday Soc. 1952, 48, 828-832.

39.

Randles, J. E. B.; Somerton, K. W., Kinetics of rapid electrode reactions. Part 3. - Electron
exchange reactions. Trans. Faraday Soc. 1952, 48, 937-950.

40.

Randles, J. E. B.; Somerton, K. W., Kinetics of rapid electrode reactions. Part 4. - Metal
ion exchange reaction at amalgam electrodes. Trans. Faraday Soc. 1952, 48, 951-955.

41.

Natan, A.; Kuritz, N.; Kronik, L., Polarizability, susceptibility and dielectric constant of
nanometer-scale molecular films. A microscopic view. Adv. Funct. Mat. 2010, 20, 20772084.

42.

Akkerman, H. B.; Naber, R. C. G.; Jongbloed, B.; van Hal, P. A.; Blom, P. W.; de Leeuw,
D. M., Electron tunneling through alkanedithiol self-assembled monolayers in large-area
molecular junctions. Proc. Nat. Acad. Sci. 2007, 104, 11161-11166.

65

43.

Pillman, H. A.; Blanchard, G. J., Molecular Consequences of Ethanol Incorporation into
Phospholipid Bilayers. J. Phys. Chem. B 2010, 114, 3840-3846.

44.

Mitchell, D. C.; Litman, B. J., Effect of Cholesterol on Molecular Order and Dynamics in
Highly Polyunsaturated Phospholipid Bilayers. Biophys. J. 1998, 75, 896-908.

66

CHAPTER 4: USING DIFFUSIONAL MOTION TO GAUGE
INTERFACIAL ADHESION OF SUPPORTED ALKYL
PHOSPHONATE MONOLAYERS

Adapted with permission from:
S. M. Baumler, Thomas M. Reidy, and G. J. Blanchard; Diffusional motion as a gauge of fluidity
and interfacial adhesion. Supported alkylphosphonate monolayers. Journal of Colloid and
Interface Science (2016) 145–155.

67

4.1

Abstract
Diffusion measurements were used to gauge the fluidity and surface binding properties of

a molecular monolayer. The monolayer film consists of octadecyl-1-phosphonic acid (ODPA) and
controlled amounts of a lyso-phosphatidic acid tagged with the fluorescent probe BODIPY
(BLPA). The monolayer films were formed using a Langmuir-Blodgett (LB) trough and deposited
onto a glass slide. Monolayer morphology was characterized during film formation using Brewster
angle microscopy (BAM). Fluorescence Recovery After Photobleaching (FRAP) microscopy was
used to measure translational diffusion of BLPA and Fluorescence Anisotropy Decay Imaging
(FADI) was used to measure rotational diffusion of the BLPA chromophore. These results provide
information on the motional freedom of the probe and, importantly, on the strength of interaction
between the probe and the support. Compositional variations in the monolayer give rise to changes
in constituent dynamics that reflect intermolecular interactions.

4.2

Introduction
Monolayer films have been examined extensively over the past half-century because of

their ability to alter the properties of an interface for a desired purpose. A single layer of molecules
can be used to change the polarity of a surface, or to mediate electron transfer, for example. With
technology requiring ever smaller dimensions, it is important to gain molecular-level control over
not only the chemical identity of interfacial layers, but also to be able to control their physical
properties.1 We have undertaken the examination of a representative interfacial monolayer with
the aim of quantitating the fluidity and adhesion of the adlayer to the interface. Understanding
interfacial dynamics is a prerequisite to controlling these properties.

68

One family of monolayer films that has received extensive research attention is selfassembled monolayers (SAMs) of alkanethiols on gold.2-13 That work has demonstrated the
delicate balance of forces required to produce a monolayer and at the same time many practical
applications have stemmed from this chemical system, including patterned interfaces for
lithographic and other applications.14-17 In addition to the alkanethiol SAMs, Langmuir-Blodgett
(LB) films have enjoyed wide use.18-21 The interactions between the amphiphilic film constituents
and the supporting surface are fundamentally different for the two systems, with physical
robustness favoring alkanethiol SAMs.12-13 LB films, however, allow for greater control over
composition and ordering, and also produce films that exhibit controllable mobility.22 This latter
feature is important especially in cases where films benefit from defect mitigation or the ability to
adapt to interfacial features. Both approaches to monolayer film formation allow complex
molecules to form well-ordered two-dimensional assemblies. For both systems, an understanding
of the interrelationship between the nanoscale heterogeneities at the support surface and defects
within the film itself, and their effect(s) on the long-range order of the films, remains to be
developed fully.
Previous work by the Talham group has shown that films of considerable ordering and
structure can be prepared using LB methodology using organophosphonic acids with divalent
metals at the appropriate subphase pH.23-24 We are concerned with the introduction of structural
heterogeneities into an amphiphilic monolayer structure, and how the film dynamics and
organization change with the amount of “defect” species present. We use octadecylphosphonic
acid (ODPA) as the amphiphile for the formation of the LB film and a fluorophore-tagged
lysophospholipid (BLPA) as the dissimilar defect molecule. Because BLPA contains the BODIPY
chromophore, it also serves as the optical probe in this work. We have chosen to use phosphonate

69

LB films in this work because, ultimately, there is greater structural versatility and range of
dynamics available with phosphonates than the corresponding carboxylates.23-29
The BLPA probe has been used to measure the diffusional properties on both short
(molecular) and long (Âľm) length scales using rotational- and translational-diffusion
measurements, respectively. Comparison of these two physical quantities affords quantitative
information on the role of the defect molecule in mediating organization within the film and on
the strength of interaction between the film and the support. While the study of dynamics in LB
and related films is not new,22, 30-34 our approach to comparing diffusional processes across a range
of length scales is both novel and information-rich. The forces binding the adlayer to the interface
were found to exceed those expected for simple physisorption, and more closely approximate the
strength of hydrogen bond enthalpies in water. The ability to quantitate such interactions opens
the door to better understanding how to control the properties of supported monolayer films and,
simultaneously, provides direct information on the molecular motion that characterizes these
interfaces.

4.3

Experimental
Materials. Octadecylphosphonic acid (ODPA) (C18H39O3P, 97% purity) (Fig. 4-1), barium

chloride dihydrate (BaCl2¡2H2O, Reagent Grade), and tetrahydrofuran (THF, 99.9%) were
obtained from Sigma Aldrich and used as received. The chromophore used in these studies was
1-(12-[4-(dipyrrometheneboron difluoride) butanoyl] amino) dodecanoyl-2-hydroxy-sn-glycero3-phosphate (ammonium salt) (BLPA) (TopFluor Lyso PA, >99%, Avanti Polar Lipids) (Fig. 41). Water (18 MΊ-cm) was obtained from a Milli-Q Plus water purification system and used for
all experiments.

70

Figure 4-1: Chemical structures of the components used in the formation of the LB film.
Octadecylphosphonic acid (ODPA), and 1-(12-[4-(dipyrrometheneboron difluoride)
butanoyl] amino) dodecanoyl-2-hydroxy-sn-glycero-3-phosphate (ammonium salt)
(BLPA).

Langmuir Monolayer Preparation and LB Film Deposition. The barium ODPA (BaODPA) LB films were prepared by the Langmuir-Blodgett technique.18-20 We used a large TeflonÂŽ
Langmuir trough (KSV Nima) equipped with a dipping mechanism and a platinum Wilhelmy plate
attached to a balance with an automated feedback system to maintain a constant surface pressure
during film transfer. Herein, we use the term ‘Langmuir monolayer’ to refer to a monolayer at the
air-water interface and LB film to refer to the monolayer transferred to the solid support.
The glass substrates were standard microscope slides (1 mm × 25 mm × 75 mm, Globe
Scientific) and were placed in a piranha solution (1H2O2:3H2SO4, Caution! Strong oxidizer!) for
30 min prior to transfer to the LB-trough for dipping. After cleaning, the substrates were rinsed
with water and placed in a covered beaker containing water until they were used for deposition.
To produce stable monolayers the subphase was a 5 mM BaCl2 aqueous solution. The subphase
pH was measured to be 5.4. The temperature of the subphase was maintained at 23° ¹ 0.1° C using
a circulating water bath (Fisher) attached to the trough. For each dipping experiment 150 ÎźL of a
1 mg/mL ODPA in THF solution was used for monolayer spreading.
71

Solutions containing 5 mol% BLPA (BLPA-5) and 10 mol% BLPA (BLPA-10) were
prepared by mixing appropriate amounts of the ODPA solution and 1 mg/mL BLPA in THF
solution and mixing for 30 sec by hand. The amount of solution spread on the surface was
calculated to give an initial surface pressure π below 1 mN/m. During spreading, care was taken
to prevent the spreading needle from touching the surface as well as preventing the surface pressure
π from rising above 0.5 mN/m in order to ensure complete spreading. After spreading, the
monolayer was allowed to equilibrate for 30 min to ensure solvent evaporation and monolayer
relaxation, before the barriers were moved inward at 5 mm/min (mean molecular area compression
of <1 Å2/min) to compress the surface monolayer to the pressure used for dipping (30 mN/m). The
compressed monolayer was allowed to equilibrate for 15 minutes before it was transferred onto
the glass slide. The monolayer was transferred (upstroke) at a dipping speed of 5 mm/min while
keeping the pressure constant, covering a total area of 35mm x 25mm on each side of the glass
slide. The coated slides were dried in air for at least 1 hr. before FRAP and FADI measurements
were made.

Brewster angle microscopy. We characterize the morphology of the monolayers formed
on the LB-trough using a Brewster angle microscope (BAM, Accurion UltraBAM).

This

instrument is characterized by 2 Âľm spatial resolution and an adjustable angle of incidence. The
laser wavelength used for BAM measurements is 653 nm (diode laser) and the instrument is
operated using software supplied by the manufacturer.

72

Figure 4-2: Idealized diagram of the prepared ODPA/BLPA Langmuir film. The films
are composed of the (a.) phosphorylated silica support, (b.) barium ion layer, and (c.)
ODPA/BLPA layer.

Steady State Fluorescence Measurements. Excitation and emission spectra of the BLPA
chromophore in THF was collected using a Jobin Yvon Spex Fluorolog-3 spectrometer. Emission
spectra were collected for BLPA in THF as well as the 5 mol% and 10 mol% samples to verify the
presence and band positions of the incorporated BLPA chromophore. Spectral resolution was set
to 1 nm for both the excitation and emission monochromators for all measurements.

73

Optical ellipsometry. Optical ellipsometry measurements were performed to determine LB
film thickness using a J. A. Woollam Co., Inc. model EC270 spectroscopic ellipsometer with a
fixed angle (75°) and wavelength range of 400-1100 nm. The EC270 is equipped with 75W Xenon
lamp source and diode array detector to simultaneously detect 44 wavelengths. The LB films were
prepared on single crystal (100) silicon wafers (University Wafer, Boston, MA). The fitting for
the ODPA layer thickness was accomplished by modeling the silicon substrate as adlayers of
silicon dioxide with fixed thickness and refractive index of 1.46, and a generic organic film layer
with a refractive index of 1.5. The silicon dioxide adlayer was determined on a freshly cleaned
substrate and measured to be 17 ± 1 Å.

Fluorescence Recovery After Photobleaching (FRAP) Measurements.

FRAP

measurements were conducted on a Nikon C2+ confocal laser scanning microscopy system
consisting of a Nikon Eclipse Ti-E inverted microscope with a confocal scanning system (Nikon
Ti-S-CON). The BLPA was excited with the 488 nm line of an Ar+ laser (Nikon Lu-N4), and
emission was detected at 525 nm using a standard PMT detector (Nikon C2-DU3). A 10x objective
lens was used for the LB films. Initial intensities (I0) were measured for a minimum of 10 s. The
bleaching time was set to 5 s, and the recovery signal after photobleaching was collected over a
minimum of 6 min. The recovery signals, Ifrap(t), were normalized using eqn. 4-1, where Ibleach is
the intensity immediately following photobleaching, I0 is the maximum intensity during the 15 s
prior to photobleaching, and Inorm(t) is the calculated normalized intensity.
I norm (t ) =

I frap (t ) − I bleach

4-1

I 0 − I bleach

A freshly prepared LB film was placed above the objective and approximately 100 ÎźL of milliQ
water was added to a random location of the film. Five measurements were made within the water
74

overlayer on each side of a slide, representing a total of 10 measurements per slide, over 6 slides.
The translational diffusion constant (DT) and percent mobility were obtained by fitting the
normalized data to the Ellenberg equation (eqn. 4-2),35
12
 
 
w2
=
I (t ) I final 1 −  2
 
  w + 4π DT t  

4-2

Ifinal represents the fluorescence intensity in the photobleached region at times greater than 5 min,
w is the diameter of the photobleached spot (10 Îźm), and DT is the translational diffusion constant
(Îźm2/s). Due to normalization method (eqn. 4-1), Ifinal is the fractional mobility of BLPA in the
LB film. The recovery time for the photobleach (τ1/2) is defined as the time it takes for the depletion
recovery to reach 50% of the final intensity (eqn. 4-3),

τ1 2 =

0.75w2
π DT

4-3

Fluorescence Anisotropy Decay Imaging (FADI) Measurements. The instrument used to
acquire fluorescence anisotropy decay images has been described previously.36 Briefly, the
instrument is based on an inverted microscope optical configuration (Nikon Eclipse Ti−U). All
images were collected using a 10x microscope objective. Time-resolved data collection is
achieved using the confocal scanning unit (Becker & Hickl DCS-120) attached to an output port
of the microscope and is controlled by a galvo-scanning drive unit (Becker & Hickl GDA- 120).
The confocal scanning unit (DSC-120) is equipped with a polarizing beam splitter and two
avalanche photodiode detectors (ID-Quantique ID100). Polarized fluorescence transients are
acquired using time-correlated single photon counting (TCSPC) detection electronics (Becker &
Hickl SPC-152, PHD-400N reference diode) and were used to determine the induced orientational
anisotropy decay function (eqn. 4-4).
75

The light source for these experiments was a synchronously pumped cavity dumped dye
laser (Coherent 702) excited by the output of a passively mode locked Nd:YVO4 laser (Spectra
Physics Vanguard). The dye laser is cavity dumped (Gooch and Housego 64389.5-SYN-9.5−1
cavity dumper driver) to control the repetition rate. The dye laser output is characterized by ca. 5
ps pulses at a repetition rate of 4 MHz (250 ns interpulse spacing). For this work the dye laser
output was set to 490 nm using Stilbene 420 dye (Exciton) and the emission collection window
was 535 Âą 30 nm, determined by a band pass filter in the confocal scanning unit. These
wavelengths were selected based on the excitation and emission spectra of the BLPA chromophore
(Fig. 4-6, A.).
Supported LB films were placed above the microscope objective and approximately 100
ÎźL of Milli-Q water was added to a random location of the film. Two measurements were made
within the hydrated region on each side of the slide, representing a total of 4 measurements per
slide over 4 slides. Each measurement produced a 256x256 pixel2 image with 256 time points
from both the parallel and perpendicular channels. The images were binned to average the data
over a 4 pixel x 4 pixel region. Anisotropy decay time constants were obtained by fitting the
induced anisotropy decay equation, which is calculated from the averaged polarized transients
(eqn. 4-4):
R (t ) =

I  (t ) − I ⊥ (t )

4-4

I  (t ) + 2 I ⊥ (t )

The anisotropy decay time constant (τHR) or can be determined by fitting R(t) to eqn. 4-5, which is
derived from the hindered rotor model.37-38
R ( t )= R ( ∞ ) + ( R ( 0 ) − R ( ∞ ) ) exp ( −t τ HR )

4-5

76

The terms R(0) and R(∞) are the zero- and infinite-time anisotropies, respectively, and the term
τHR is the anisotropy decay time constant. We consider the physical significance of these data
below.

4.4

Results and Discussion
We are concerned with the fundamental properties of LB films. These include the stability

and fluidity of the film as a function of how it was formed, and the role that probe molecules may
play in mediating film organization and dynamics. We use Langmuir isotherms (π/A) to examine
changes in the monolayer density as a function of applied pressure, ionic strength of the subphase,
and the addition of selected amounts of BLPA. We use a suite of optical techniques to evaluate
the morphology (BAM) and dynamics (FADI, FRAP) of the LB films as a function of system
conditions. We consider the information content of these data below, focusing first on the role of
subphase composition on Langmuir monolayer properties.

Effect of subphase ionic strength on ODPA Langmuir monolayer morphology. Surfacepressure vs. area isotherms (π/A-isotherms) have been acquired for monolayers prepared over an
aqueous subphase containing no ionic constituents and a subphase containing 5 mM BaCl2 (Fig.
4-3, above). BAM images were collected at 5mN/m surface pressure increments for each sample.
The isotherm for the monolayer formed on the water subphase consists of a sharp increase in
pressure at a mean molecular area of 21 Å2 and a characteristic collapse of the monolayer at ca.
62 mN/m. For the subphase containing 5 mM BaCl2, the π/A-isotherm takes a different form. The
isotherm does not begin to exhibit a significant increase in pressure until ca. 18 Å2/molecule, has
a mean molecular area of 13 Å2, and the slope is more shallow (11 mN/m/Å2) compared to the

77

isotherm of the monolayer formed on the water subphase (16 mN/m/Å2). Additionally, the Ba2+containing monolayer does not exhibit a collapse at high pressure similar to the monolayer formed
on the water subphase. Rather, the monolayer formed on the BaCl2-containing subphase maintains
a constant pressure at ca. 60 mN/m with decreasing surface area. BAM images of the two
monolayers reveal differences beginning at formation, before any compression occurs (Figs. 4-3
A, D). The monolayer formed on the water surface exhibits fluid regions with small domain
ordering present as the white elliptical/circular regions with smaller holes throughout. The
monolayer formed with BaCl2 in the subphase exhibits larger island-like regions with dark domain
boundaries. Upon compression of the monolayer on the water subphase, the images appear
uniform with small darker lines throughout, up to the point of collapse, when bright lines and
cracks begin to appear (Figs. 4-3, C, F). The monolayer formed on the subphase containing BaCl2
exhibits scattered bright islands, which do not appear to change in size and/or shape upon
compression; these island regions appear to reorganize to fill in the liquid regions with increasing
pressure. The Talham group has shown previously that the formation of crystalline Ba/ODPA
monolayers is highly pH dependent, and can be formed around pH 8-9.23 We attribute much of
the heterogeneity within these BAM images to be due the subphase pH being lower than the
optimal pH for deposition. Ellipsometric thickness measurements after deposition onto the glass
slides confirm thicknesses consistent with a monolayer (27 ± 1 Å).29

78

Figure 4-3: Above: 𝜋𝜋/A-isotherm of ODPA on an aqueous subphase (red solid) and with a
5 mM Ba2+ subphase (blue dash). 𝜋𝜋 /A-isotherm of BLPA on an aqueous subphase (red
dot) and with a 5 mM Ba2+ subphase (blue dot). The temperature for the subphase was
23 °C and pH = 5.4. The monolayers were compressed at a rate of 0.8 Å2/min. Orange
circles represent pressures at which BAM images (A–F) were taken. Below: Images A-C
are of ODPA on an aqueous subphase. Images D-F are of ODPA on an aqueous
subphase containing 5 mM BaCl2. Scale bars represent 100 Îźm.

79

The introduction of ionic character into the subphase allows for two phenomena to occur.
Increasing ionic strength is known to stabilize the condensed phase allowing for the crystal
structure of the long-chain aliphatic acids to be maintained during transfer to the glass support.39
Through charge compensation, the Ba2+ cation would allow the negatively charged polar
phosphonic acid headgroups to pack more closely than they would in water at a pH above the
pKa(s) of the headgroup. Due to the change in the headgroup spacing, it is expected that the tilt
angle of the aliphatic chains will be different for deposition on the two subphases,40 and our
experimental findings are consistent with this expectation.

Effect of Increasing BLPA Concentration on ODPA Langmuir monolayer Morphology.
π/A-isotherms have also been acquired for monolayers prepared over an aqueous subphase
containing 5 mM BaCl2 at two different BLPA concentrations (Fig. 4-4, Above). BLPA-5
increases the mean molecular area of pure ODPA from 13 Å2 to 25 Å2. For BLPA-10 the mean
molecular area decreases from 25 Å2 for BLPA-5 to 17 Å2. The slopes of the isotherm curves
increase with increasing BLPA concentration, from 11 mN/m/Å2 for pure ODPA, to 13 mN/m/Å2
for BLPA-5, and 20 mN/m/Å2 for BLPA-10. The decrease in mean molecular area and increase
in slope correlated with the increase in BLPA concentration can be explained in the context of an
increased packing density of the BLPA through phase segregation.

80

Figure 4-4: Above: 𝜋𝜋/A-isotherm of the two component monolayers containing ODPA
and BLPA on an aqueous subphase with a Ba2+ concentration of 5 mM, and 0 mol%
BLPA (red solid), 5 mol% (blue dash), and 10 mol% (green dot). The temperature for
the subphase was 23 °C and pH = 5.4. The monolayers were compressed at a rate of 0.8
Å2/min. Orange circles represent pressures at which BAM images (A-F) were taken.
Below: Images A-C contain ODPA and 5 mol% BLPA, images D-F contain ODPA and
10 mol% BLPA. Images at 0 mol% BLPA are presented in Fig. 4-3 (D-F). Scale bars
represent 100 Îźm.

81

The mean molecular areas derived from the data shown in Fig. 4-3 were found to be lower
than those reported previously.23 We attribute the rise of the isotherms at values lower than
expected to several possible causes. The first is due to the solvent system used during application
of the monolayer. ODPA was found to dissolve in THF at concentrations as high as 1 mg/mL and
BLPA also exhibited the requisite solubility in this solvent. As a consequence of this solvent
choice, the miscibility of the subphase with THF gave rise to the loss of a finite amount of material
at the subphase surface during deposition, leading to lower than expected mean molecular areas
(Fig. 4-5).

Figure 4-5: π/A-isotherm of the two component monolayers containing ODPA and
BLPA on an aqueous subphase with a Ba2+ concentration of 5 mM, and 0 mol% BLPA
(100% ODPA) (red dash), 5 mol% (blue dot), and 10 mol% (blue dash-dot) dissolved in
THF when applied to trough. The application of ODPA when dissolved in chloroform at
0.2 mg/mL and added under similar conditions is shown (red solid).
82

The second possible reason is due to the solubility of the BLPA chromophore in the subphase at
increasing pressures (Fig. 4-3). When comparing the pure BLPA on water to pure BLPA on the
barium chloride sub-phase, the addition of ionic character improves the formation of a stable
monolayer slightly, likely by decreasing the solubility of the BLPA (Fig. 4-3).
BAM images of the monolayer without BLPA formed with BaCl2 in the subphase are
shown in Fig. 4-4. The addition of 5% BLPA creates two distinct regions (Figs. 4-4, A, D). Upon
compression, both BLPA-5 and BLPA-10 exhibit bright cracks at the collapse pressure (Figs. 4-4,
C, F). BLPA-5 contains ca. 50% more bright regions than BLPA-10, and the bright regions of the
BLPA-5 are, on average, ca. 20% smaller than those in BLPA-10. We believe these bright spots
are aggregates of BLPA dispersed throughout the monolayer and the differences in quantity of
bright regions correlates well with the two-fold increase in amount of BLPA present.

Formation of Aggregates at Higher BLPA Concentrations. The steady state absorbance
and emission spectra of BLPA in THF, as well as the emission for BLPA-5 and BLPA-10 LB films
are shown in Fig. 4-6, A. Increasing the concentration of BLPA in the LB films causes a
bathochromic shift of the emission spectrum of ~6 nm. The difference in band shift between the
BLPA-5 and BLPA-10 films is ~4 nm.
The BODIPY chromophore is known to form either H- or J- aggregates,41-44 depending on
the environment and functionalities attached to the BODIPY chromophore.45-48 We attribute the
slight bathochromic shift to the formation of J-aggregates in these films. The peak shift, whether
bathochromic or hypsochromic, depends on the angle between the constituent chromophore
transition dipole moments (Fig. 4-6, B). J-aggregation is characterized by the angle between the
constituent transition moments, γ ≤ 54.7°. As the angle between the BODIPY transition moments

83

approaches zero, the J-aggregates are characterized by a head-to-tail orientation, while Haggregates are characterized by a transition moment angle γ > 54.7°, where the transition dipole
moments are stacked face-to-face or there is considerable π-π stacking in the aggregate. We
attribute the observed spectral shift to aggregation, with the increased disorder induced by the
BLPA-10 films decreasing the angle between the two chromophores and allowing for a more headto-tail configuration.

Figure 4-6: (A.) Emission spectra of the two component LB films. BLPA-5 (blue solid),
and BLPA-10 (green dash). Absorbance and emission spectra of BLPA in THF (black
dot). All spectra were normalized to unity. Schematic representation of a structural
description of H- and J-aggregates of the BODIPY chromophore (B.) Steady state
fluorescence images of BLPA-5 are shown in (C.) and BLPA-10 in (D.). Scale bars
represent 500 Îźm.
Steady state fluorescence images of BLPA-5 and BLPA-10 are shown in figure 4-6, C and
4-6, D, respectively. Increasing the concentration of BLPA in the LB films reveals an increase in
84

pinhole defects and increased heterogeneity, evidenced by the bright striped regions the BLPA-10
image (Fig. 4-6, D).

The BLPA-5 image shows a considerable number of bright regions

throughout the image. We attribute these bright spots to regions of BLPA aggregation. There is
considerably less intense spotting in the BLPA-10 image. We attribute the decrease in spotting
intensity to be due to two main factors. First, at higher concentrations the contrast between the
BLPA aggregates and the larger film area decreases due to the overall increased presence of BLPA
chromophore. Second, BLPA-5 shows larger regions of uniform intensity compared to BLPA-10
with alternating regions of high and low intensity (striping). This observation suggests not only
that more BLPA is present, but BLPA may phase segregate into regions where the film exhibits
different domain organization.

Translational Diffusion Slowed by Aggregation at Increased BLPA Concentrations.
Macroscopic translational diffusion of the ODPA/BLPA LB films was measured using FRAP.
FRAP is sensitive to the organization of the LB film. FRAP data for both BLPA-5 and BLPA-10
films are shown in Fig. 4-7 and Table 4-1.
Table 4-1: Translational diffusion constants (DT), Mobile fraction %, and Recovery
speed (τ1/2) determined for the 5 mol% and 10 mol% substrates from FRAP
measurements.
Molar % BLPA
Mobile Fraction (%) DTeff (Îźm2/s)
τ1/2 (s)
5%
16 Âą 4
0.26 Âą 0.09
90 Âą 30
10%
23 Âą 4
0.09 Âą 0.02
250 Âą 50
Errors are ¹ 1σ for a minimum of 5 individual determinations on each substrate, over a minimum
of 6 substrates. 5% and 10% values were determined to be significantly different to 95%
confidence.
The average translational diffusion constant of the BLPA-5 film is 0.26Âą0.09 Îźm2/s with a mobile
fraction of 16Âą4 %. The BLPA-10 film gives DT = 0.09Âą0.02 Îźm2/s with a mobile fraction of
23Âą4 %. The time constant for recovery of the BLPA-5 film is 90Âą30 s and for the BLPA-10 film

85

τ = 250±50 s. These data were determined from the raw time-scans by fitting to Eqs. 4-2 and 4-3
. The three-fold decrease in both DT and τ indicate that translational motion of BLPA-10 is more
hindered, and we assert that the reason for this is the aggregation of chromophores (vide infra).
We consider the energetic implications of these findings below.

Figure 4-7: FRAP recovery of the BLPA chromophore in (A.) BLPA-5 (red) and (B.)
BLPA-10 (blue). The green lines represent best-fit lines using the Ellenberg equation
with residuals plotted below. (C.) Comparison between the BLPA-5 and BLPA-10
curves is shown. All FRAP curves were normalized prior to fit.

Rotational Diffusion and Information Content. FADI measurements have been performed
for both BLPA-5 and BLPA-10 LB films (Table 4-2). Typical fluorescence anisotropy decays for
the films are shown in Fig. 4-8. To gain greater insight into the effect of increased BLPA
concentration on the surface structure and dynamics, we

86

Figure 4-8: Typical experimental anisotropy decay functions for the BLPA-5 (A.) and
BLPA-10 (D.) LB films with residuals of best fit inset are shown. Fluorescence transients
were fit to determine the fluorescence lifetime shown in (B., BLPA-5) and (E., BLPA-10).
The colors shown represent the lifetime value at a given pixel, with blue being short
lifetimes and red being longer lifetimes. Fluorescence anisotropy decay time constant
images are shown in (C., BLPA-5) and (F. BLPA-10). The colors shown are set by the
imaging software, with blue being short decay time constants and red being longer decay
time constants. All images were acquired using a 10x microscope objective. Scale bars
represent 500 Îźm.

87

have acquired 256x256 pixel images of the fluorescence lifetime and anisotropy decay of the films
(Figs. 4-8, B-C, E-F). These data provide qualitative insight into how structural heterogeneity is
induced at higher BLPA concentrations. BLPA-10 films have characteristically shorter average
fluorescence lifetimes and longer anisotropy decay time constants. The shorter chromophore
lifetime for the BLPA-10 films suggests quenching. We assert that the fluorescence quenching is
due to the formation of BODIPY aggregates, consistent with the steady-state emission shift
reported for these films (vide supra). The anisotropy decay images for the BLPA-10 films show
both a broader distribution and longer anisotropy decay time constants than are seen for the BLPA5 films. These data are consistent with chromophore aggregation at higher concentration.
Table 4-2: Reorientation time (𝜏𝜏HR), rotational diffusion constant (DROT), and rotational
wobbling constant (DW) determined by the local Environment of BLPA in a Ba-ODPA
LB film.
Molar % BLPA
τHR (ps)
DROT (MHz)
Dw (MHz)
5%
3,400 Âą 220
49.0 Âą 3.1
30.9 Âą 2.0
10%
4,000 Âą 200
41.8 Âą 2.1
26.5 Âą 1.3
Errors are ¹ 1σ for a minimum of 2 individual scans over an area of Οm2 on each substrate, over
a minimum of 4 substrates. 5% and 10% rotational values were determined to be significantly
different to 90% confidence.
To obtain quantitative information on the chromophore local environment, the images were
analyzed as described above to obtain the average anisotropy decay time constant by fitting the
collected fluorescence anisotropy decay transients to eqn. 4-5. The reorientation times for the
films were τHR = 3,400 ± 220 ps and 4,000 ± 200 ps for the BLPA-5 and BLPA-10 films,
respectively. The LB film under investigation here contains a tethered chromophore, so the
hindered rotor model is appropriate (Eqs. 4-5 and 4-6).37-38

τ HR

7θ 02
=
24 DW

12
12
 




∞
R
(
)
−1 
=
+ 1 − 0.5 
θ 0 cos 0.5 8 
  R ( 0 ) 


  






4-6

88

In this model, the (fast) chromophore motion is constrained by the medium surrounding it
and its tether to the support surface. Rotational motion can be described as occurring in a cone of
semi-angle θ0, and there will be a diffusive “wobbling” of the chromophore about its tethering
bond. This motion is characterized by the quantity Dw. The relationship between Dw and DROT
can be estimated by the approximation that the experimental τ ~ τOR ~ τHR,

τ ≈ τ OR ≈ τ HR=
DROT

7θ 02
1
=
6 DROT 24 DW

4-7

 4 
=  2  DW
 7θ 0 

The rotational diffusion constants were determined to be DROT = 49.0 Âą 3.1 MHz and DROT = 41.8
Âą 2.1 MHz for the BLPA-5 and BLPA-10 films, respectively (eqn. 4-7). The slower time constant
in BLPA-10 is consistent with aggregation of the chromophores.
As can be seen in eqn. 4-7, the relationship between DROT and Dw depends on the angle of
the confining cone (Fig. 4-9, A.). For θ0 = 0, the relationship is undefined, and this situation
corresponds physically to the probe being immobile and unable to exhibit diffusional motion. For
physically realistic angles, between ca. 30° and 90°, the coefficient (4/7θ02) varies between 2.08

Figure 4-9: Schematic of equilibrium system under consideration.
89

and 0.23, for θ0 = 54.7°, the magic angle, 0.63, and for θ0 = 43.4°, DROT = Dw. Owing to the
heterogeneity in both the BLPA-5 and BLPA-10 films, the BLPA probe population is likely
characterized by a random orientational distribution. For such a distribution the confining cone
semi-angle distribution averages to 54.7°,37-38 yielding a value of DROT = 0.63Dw = 30.9 ¹ 2.0 MHz
for BLPA-5, and 26.5 Âą 1.3 MHz for BLPA-10.

Implications of the Ba-ODPA/BLPA system thermodynamics on the diffusional dynamics.
Comparing rotational diffusion and translational diffusion data on the same system can provide
substantial insight into the properties of the system, with the detailed information content
depending on the system. As noted above, chromophore motion is described in the context of the
hindered rotor model,37-38 and it is the relationship of the wobbling diffusion constant, Dw, for
chromophore motion about its tethering bond that we must relate to the rotational diffusion
constant, DROT. Longer range, translational diffusion of the same chromophore within the LB film
will be mediated by interactions between the amphiphile phosphonate headgroups and the support
surface, and both of these issues have to be taken into account in extracting information from these
data.
The relationship between rotational diffusion and translational diffusion is understood for
a spherical particle is a uniform medium, where the rotational diffusion constant is given by the
Debye-Stokes-Einstein equation (eqn. 4-8) and the translational diffusion constant is given by the
Stokes-Einstein-Sutherland equation (eqn. 4-9).
DROT =

k BT
8πη r 3

4-8

DROT =

k BT
6πη r

4-9

90

where Ρ is the viscosity of the medium, r is the radius of the diffusing particle and kBT is the
thermal energy term. There are well-established correction factors for eqn. 4-8 to account for the
boundary condition between the particle and the medium in which it is immersed, as well as for
the non-spherical shape of the particle, but those correction factors yield only small changes from
eqn. 4-8 for the systems of interest, and their inclusion does not bear on the fundamental
relationship between DROT and DT (eqn. 4-10).

 4r 2 
DT = 
 DROT
 3 

4-10

Eqn. 4-10 is frequently not obtained experimentally because of the specific physical and chemical
conditions of the system under investigation.
For the LB films reported here the amphiphile constituents interact with the solid support
and it is possible to estimate the energy of that interaction. Experimentally, the value of DT
recovered, DTeff, is less than that expected if the adlayer were not interacting with the support (eqn.
4-9). There are a variety of ways to interpret a value of DTeff less than that predicted by eqn. 4-9.
While heterogeneities in the films certainly exist, including pinholes, regions of disproportionate
ODPA:BLPA concentrations due to aggregation, or the potential formation of multilayer islands,
the DT and DROT values are acquired as average values over tens of microns of film surface. For a
heterogeneous system such anomalous diffusion is considered to result from frustrated
chromophore motion.49-51 The film under consideration here contains only one amphiphile in
addition to the chromophore, therefore the notion of anomalous diffusion does not apply. For a
system where there is an equilibrium between the free and surface-bound amphiphile, the value of
DTeff reflects the fraction of time the chromophores are free to diffuse, i.e. not bound to the support.
In this treatment we focus on the dynamic rather than the steady state (mobile fraction) contribution
to the FRAP data. The steady state contributions to these data appear to be the same for the two
91

chromophore concentrations to within the experimental uncertainty (Table 4-1), and the
normalization of the experimental FRAP data can, in principle, influence the recovered value for
the mobile fraction, where time-domain data are not affected by normalization.
We model this equilibrium according to eqn. 4-11, with the diffusion constant of the bound
species being zero.
DTeff =
Afree DT + Abound DTbound =
Afree DT
Afree + Abound =
1

4-11

DTbound = 0

The free fraction, Afree, is related to the equilibrium constant for this system (Fig. 4-9, B.). If the
bound amphiphiles have a surface coverage φ, then the free sites are given by 1-φ, which is equal
to Afree, and the equilibrium constant for this system is

k free
=
K eq =
kbound

(1 − φ )

2

4-12

φ

The fraction of free sites, Afree, is related to Keq by

 ( 2 + K ) − K 2 + 4K
eq
eq
eq
Afree = 1 − 

2







4-13

Thus determination of Afree from the experimentally-derived quantities Dw, θ0 and DTeff provides
Keq.
We calculate the hydrodynamic volume of BLPA to be 602 Å3, corresponding to r = 5.2 Å
for a spherical diffusing species. Using eqn. 4-10 we calculate DT = 11.1 Âą 0.7 Âľm2/s for BLPA5 and DT = 9.6 Âą 0.5 Âľm2/s for BLPA-10 using DROT values derived from the experimental FADI
data.

92

The experimental values of DTeff for BLPA, determined by FRAP, are 0.26 Âą 0.09 Âľm2/s
for BLPA-5 and 0.09 Âą 0.02 Âľm2/s for BLPA-10. Using eqn. 4-11 we determine Afree = 0.022 Âą
0.009 for BLPA-5 and 0.009 ± 0.002 for BLPA-10. From Afree = 1-φ, we determine Keq = 5.0
(+4.9,-3.2) x10-4 for BLPA-5 and 8.2 (+4,-3.3) x10-5 for BLPA-10. As the equilibrium constant
for this system is cast as a dissociation constant, we can calculate the free energy of dissociation
for the film constituents (Table 4-3).
Table 4-3: Calculated Diffusion constant (DT), Unbound free fraction (Afree), Dissociation
equilibrium constant (Keq), and Free energy for dissociation (ΔGdis) for the BLPA in a
Ba-ODPA LB film.
Mole% BLPA
Keq (x10-4)
ΔGdis
DT0 (Âľm2/s) Unbound
fraction (Afree)
(kJ/mol)
5%
11.1 Âą 0.7
0.023 Âą 0.009
5.0 (+4.9,-3.2)
18.5 (+2.6,-1.7)
10%
9.6 Âą 0.5
0.009 Âą 0.002
0.8 (+0.4,-0.3)
22.9 (+1.3, -0.9)
∆Gdis =
− RT ln K eq

4-14

For BLPA-5 we find ∆Gdis = 18.5 (+2.6,-1.7) kJ/mol and for BLPA-10, ∆Gdis = 22.9 (+1.3,
-0.9) kJ/mol. There are several important pieces of information contained in these data. The first
important point to note is that (dipolar) aggregation is more prominent for BLPA-10 than for
BLPA-5, and the effective change in the net driving force for adlayer formation due to an unknown
amount of aggregation is energetically favorable by ca. 4 kJ/mol. The second is that the strength
of interaction between the phosphate amphiphile and the Ba/SiOx surface is very similar to that of
the hydrogen bond enthalpies of water (ca. 23 kJ/mol),52 and consistent with previous results,53
but somewhat larger than that reported for water in proximity of nonpolar species.54 We attribute
much of the observed interactions to be due to intermolecular hydrogen bonding between ODPA
molecules or ODPA and the hydroxylated silica surface. The metal ions that were carried from
the LB trough subphase may be facilitating interactions between the amphiphile and the support
but these interactions cannot be considered complexation in any sense.55-57 In fact, it is not likely

93

that much interaction occurs between the Ba2+ and ODPA due to the sub-optimal pH during
transfer to the support, a condition that reduces transfer of the metal ion to the support.23

4.5

Conclusions
We have investigated the dynamics of a tethered BODIPY chromophore contained in a LB

film. Our goal in this work was to quantitate the fluidity and adhesion of the adlayer to the
interface. By examining the diffusional behavior of this tethered chromophore over length scales
ranging from molecular through macroscopic, we have demonstrated a direct and novel means for
the evaluation of adlayer binding strength on a support surface. This is the first report that we are
aware of that utilizes complementary molecular diffusion measurements for this purpose.
Simultaneously, we are able to ascertain the motional freedom that is characteristic of this
chromophore and the effect of chromophore aggregation on molecular motion. These data provide
insight into the binding strength of a barium phosphonate species to a silica surface and, by
extension, provide an estimate of the binding strength for related interfaces. The value obtained
for ∆Gdis, ca. 20 kJ/mol, is entirely consistent with interactions dominated by H-bonding58 and
there is also a chromophore concentration dependence that suggests some information on the
interaction energy of aggregated chromophores.
We note that the strength of interaction for our monolayer with a silica support is very
similar to the binding interactions responsible for the formation of alkanethiol monolayers on gold
and mercury.12-13, 59 This is an important comparison to make because the spectroscopic evaluation
of chromophore dynamics in an alkanethiol/gold SAM is limited by chromophore quenching,60
and these data offer a way to at least estimate the expected dynamics for this family of systems.

94

The next step in this work is to evaluate the role of chemical modification of the support as a means
of controlling binding strength.

95

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96

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59.

Blomgren, E.; Bockris, J. O. M.; Jesch, C., The adsorption of butyl, phenyl and naphthyl
compounds at the interface mercury-aqueous acid solution. J. Phys. Chem. 1961, 65 (11),
2000-2010.

60.

Karpovich, D. S.; Blanchard, G. J., Dynamics of a tethered chromophore imbedded in a
self-assembled monolayer. Langmuir 1996, 12 (23), 5522-5524.

101

CHAPTER 5: THE INFLUENCE OF METAL IONS ON THE
DYNAMICS OF SUPPORTED PHOSPHOLIPID LANGMUIR
FILMS

Adapted with permission from:
Stephen. M. Baumler, and Gary J. Blanchard; The Influence of Metal Ions on the Dynamics of
Supported Phospholipid Langmuir Films. Langmuir (2017), 33, 2986–2992.

102

5.1

Abstract
The translational diffusion dynamics of the zwitterionic lipid, 1,2-dimyristoyl-sn-glycero-

3-phosphocholine (DMPC) at a planar phosphorylated support surface containing metal ions
(Mg2+, Ca2+, Ba2+, Ni2+, Zn2+, Cd2+, Zr4+) was investigated using X-ray photoelectron spectroscopy
(XPS) and fluorescence recovery after photobleaching (FRAP). Fluorescence recovery curves
yielded diffusion constants on the order of 2-5 Îźm2/s for the chromophore-tagged 12:0 NBD-LysoPC. Ionic interactions between the zwitterionic head-group and metal ions were found to play a
secondary role in mediating lipid fluidity. This work provides quantitative insight into the extent
to which the fluidity of a supported lipid film is mediated by the ionic interactions between headgroup and surface vs. that of the lipid-lipid tail-group interactions.

5.2

Introduction
Supported lipid monolayers are an important structural motif that can be used to understand

the fundamental processes that account for the properties of certain biomimetic interfaces and
plasma membrane structures. The monolayer structure provides a comparatively simple structural
motif for the characterization of acyl chain interactions as well as interactions between the support
and the lipid. The most common methods used to prepare supported lipid systems involve a
Langmuir approach (Langmuir-Blodgett1-2 or Langmuir-Schaeffer3) or by the rupture and
spreading of small lipid vesicles (vesicle fusion).4 Regardless of preparation technique, these
systems yield a film that maintains the diffusional properties comparable to multi-bilayer systems
and these properties are known to be dependent on the supporting surface.5
Recent studies have examined the extent to which the underlying support influences the
formation and adhesion of supported lipid bilayers prepared by the fusion deposition of small

103

unilamellar vesicles (SUVs) to a variety of supporting materials such as silica, glass, mica, various
metal oxides such as TiO2 and SrTiO3, and more recently inorganic nanoparticles.6-10 In all studies,
the underlying support was shown to have a marked effect on both the rupture and spreading
processes of the SUVs.
Similarly, both pH and ionic strength have been shown to play important roles in the
formation of supported lipid bilayers through the SUV approach.11
examined the influence of metal ions such as Mg+2,12 Ca2+,

13-16

Studies have recently

and Sr2+

17-18

on the roles of

adhesion and stability. In these studies, the metal ions were shown to provide added adhesion and
stability through the screening of charges, modifying ionic interactions between lipids and support.
In a similar fashion, pH has been shown to modulate the formation of different lipid structural
motifs through charge mediation at the metal oxide interface.19
Utilizing the formation and adhesion of SUVs, we have shown previously that robust airstable lipid layers can be formed through the aid of zirconium-phosphate (ZP) chemistry and is
dependent upon the lipid head-group used in self-assembly process.20 The DMPC lipid adlayers
formed by this approach were found to be structurally complete and nominally homogeneous
monolayers. Similar studies have examined the role to which different metal ions play in
mediating lipid-surface interactions in the formation of the lipid-metal-phosphonate films.21 The
formation of structurally robust lipid films through this lipid-metal coordination process were
found to rely upon the use of metal ions with high charge and small ionic radius. These films,
compared to their ZP counterpart, were found to be spatially heterogeneous with only partial
coverage.
While there have been many studies on the effect of metal cations, pH, ionic strength, and
supporting metal oxide surface on the formation of supported lipid films, relatively little work has

104

been done examining how these films behave after formation. Pretreatment effects, namely
changes in surface roughness, have been shown to change the diffusional properties of the
supporting lipid layers.22-23 The diffusional properties appear to be mediated, in part, by the
presence of an ill-defined water underlayer (~20-40Å) between the lipid head-groups and support
surface.24-25 Recent work has suggested that electrostatically induced ordering of the water
underlayer may alter diffusional properties of supported lipid layers due to a frictional component
from “hydration forces” between the support surface and lipid head-groups.26-27
While the presence of divalent cations has been shown to affect both the formation and
adhesion of SUVs to the surface, there are two issues that remain unresolved. The lipid interlayer
interactions can serve to mask the interactions of primary concern here, namely the support-lipid
interaction, and the presence and influence of metal ions in the water layer between the support
and the lipid remains to be elucidated fully. In the present study we have examined the influence
of not only the underlying surface, but also metal ions contained in the galley between the
supporting surface and lipid head-groups on the diffusional properties of the supported lipid
monolayer. We use the Langmuir-Blodgett method1-2 to provide a controlled means of forming
these films in the presence of the various metal ions. Diffusional measurements of chromophores
in the supported film are an effective means of characterizing interactions between the lipid and
the supporting surface as well as intermolecular interactions between the supported lipids.

5.3

Experimental
Materials. 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC) and 1-(12-[(7-nitro-2-

1,3-benzoxadiazol-4-yl)amino]dodecanoyl)-2-hydroxy-sn-glycero-3-phosphocholine

(NBD-

LPC) were purchased from AvantiÂŽ Polar Lipids, Inc. (Fig. 5-1). Chloroform was purchased from

105

Macron Fine Chemicals and used as received. 2,4,6 Trimethyl pyridine (99%), acetonitrile
(HPLC), barium chloride dihydrate (99.9%), cadmium chloride (99.9%), calcium chloride
dihydrate (99%), magnesium chloride hexahydrate (98%), nickel chloride hexahydrate (reagent
plus), phosphorus oxychloride (99%), TrizmaÂŽ Hydrochloride (>99.0%), TrizmaÂŽ base (>99.9%),
zinc chloride (>98%), and zirconium oxychloride octahydrate (98%) were purchased from Sigma
Aldrich and used as received. Water was obtained from a Milli-Q Plus water purification system
and used for all experiments.

Figure 5-1: Chemical structures of 1,2-dimyristoyl-sn-glycero-3-phosphocholine
(DMPC) and 1-(12-[(7-nitro-2-1,3-benzoxadiazol-4-yl)amino]dodecanoyl)-2-hydroxy-snglycero-3-phosphocholine (NBD-LPC).

Substrate Phosphorylation. The round coverglass substrates were composed of D263M
borosilicate glass (#1.5 x 25 mm dia., Warner Instruments) and were placed in piranha solution
(1H2O2:3H2SO4, Caution! Strong oxidizer!) for 30 min. After cleaning, the substrates were
sonicated for 5 minutes with water and thoroughly rinsed with dry acetonitrile. The substrates
were then exposed to phosphorus oxychloride (0.4 mL) and 2,4,6-trimethylpyridine (0.4 mL) in

106

10 mL dry acetonitrile for 3 hrs. The substrates were rinsed thoroughly with dry acetonitrile and
kept covered in water until needed for deposition.

Formation of DMPC Monolayers. We use the term “DMPC monolayer” in reference to a
monolayer at the air–water interface and “DMPC film” in reference to the monolayer transferred
to the solid support. Phospholipid monolayers were prepared on a Langmuir trough (Nima)
equipped with a paper Wilhelmy plate. The aqueous subphase was composed of 50 mM Tris
buffer at pH 7.5 and 1mM of the chosen metal chloride. The temperature of the subphase was
maintained at 20.0 ¹ 0.5 °C using a circulating water bath (Fisher) attached to the trough. A 30
ÎźL aliquot of a 1 mg/mL DMPC/NBD-LPC (2 mol%) in chloroform solution was used for
monolayer spreading. During spreading, care was taken to prevent the spreading needle from
touching the surface as well as preventing the surface pressure from rising above 0.5 mN/m in
order to ensure complete spreading. After spreading, the monolayer was allowed to equilibrate for
15 min. to ensure solvent evaporation and monolayer relaxation. The barriers were moved inward
at a rate of 5 mm/min (<1 Å2/molecule-min) until monolayer collapse. The isotherms were equally
adjusted to account for solvent evaporation in the DMPC/NBD-LPC solution by 5 ÎźL/hr.

Formation of Phospholipid Films.

The phospholipid films were prepared by the

Langmuir–Blodgett technique.1-2 A large Teflon® Langmuir trough (KSV Nima) equipped with a
dipping mechanism and a platinum Wilhelmy plate attached to a balance with an automated
feedback system was used to maintain a constant surface pressure during film transfer. The
temperature of the subphase was maintained at 20.0 ¹ 0.5 °C using a circulating water bath (Fisher)
attached to the trough. The subphase and monolayer spreading solution were as described above.

107

The larger trough required 70 ÎźL of a 1 mg/mL DMPC/NBD-LPC (2 mol%) solution. The barriers
were moved inward at a rate of 5 mm/min (<1 Å2/molecule-min) to compress the surface
monolayer to 25 mN/m. The compressed monolayer was allowed to equilibrate for 15 min before
it was transferred onto the glass support. The monolayer was transferred (upstroke) at a dipping
speed of 5 mm/min while keeping the pressure constant. Transfer ratios for the films were
measured to be 1.0 Âą 0.1. The coated slides were placed in an AttofluorÂŽ cell chamber (Invitrogen)
and gently covered with the subphase solution before FRAP measurements were made.

Fluorescence Recovery After Photobleaching (FRAP).

FRAP measurements were

conducted on a Nikon C2+ confocal laser scanning microscopy system consisting of a Nikon
Eclipse Ti-E inverted microscope with a confocal scanning system (Nikon Ti-S-CON). The NBDLPC was excited with the 488 nm line of a diode laser (Nikon Lu-N4), and emission was detected
at 525 nm using a standard PMT detector (Nikon C2-DU3). A 20x objective lens was used for the
films.

Two independently prepared DMPC films were measured with a minimum of 10

measurements at random locations. Initial intensities (Ipre) were measured for 10 s. The bleaching
time was set to 2 s at 100% laser power, and the recovery signal Ifrap(t) after photobleaching was
collected over 2 min.
To account for acquisition bleaching, the recovery signals, Ifrap(t), were normalized using
double normalization described by Phair et al. (eqn. 5-1).28 Ifrap-pre is the average stimulation region
intensity prior to photobleaching, Iref-pre is the average reference intensity prior to photobleaching,
Iref(t) is reference intensity after photobleaching, and frap(t) is the calculated normalized intensity.
=
frap
(t )

I ref − pre I frap (t )
⋅
I ref (t ) I frap − pre

5-1

108

XPS Measurements. XPS measurements were performed on a Perkin-Elmer Phi 5600
ESCA instrument equipped with an Al Κɑ X-ray source to illuminate the sample at a take-off angle
of 45°. Samples were analyzed at pressures between 10-9 and 10-8 Torr with a spot size is ca. 250
mm2 and a pass energy of 187.9 eV and 29.4 eV for the survey and detailed scans, respectively.
Survey scans of 0–1100 eV binding energy and detailed scans for the metal ion of interest as well
as C1s, O1s, P2p, and Sn 3d5/2 or Si2p were measured for all samples. All peaks were corrected
in reference to the C1s peak associated with adventitious C at 284.8 eV. Peak areas were
determined by Shirley background subtraction and an area correction using calibrated sensitivity
factors.

5.4

Results and Discussion
Langmuir monolayer organization mediation by metal ion identity. To better understand

the influence of metal ion identity on the dynamics and physical characteristics of a supported
zwitterionic phospholipid monolayer, surface pressure vs. mean molecular area isotherms were
performed (Figs. 5-2 and 5-3). Changes in the isotherm were examined vs. changes in the metal
ion identity.

109

Figure 5-2: Surface pressure / mean molecular area-isotherm of the DMPC/NBD-LPC
monolayers formed on subphases containing buffer (red solid), and 1 mM MgCl2 (blue
dash), CaCl2 (green dot), or BaCl2 (purple dash-dot). Inset: Expanded region showing
mean molecular area at dipping pressure.
We first examined the effect of the alkaline earth metals (Mg2+, Ca2+, Ba2+) on the
isotherms. The alkaline metals showed an increase in mean molecular area vs. DMPC on the
buffer phase containing no metal ions (we discuss the implications for this increase below).
However, there was only a slight discernable difference between the alkaline earth metal identity
and the mean molecular area of the DMPC monolayer. Next we examined the effect of a series of
transition metals (Ni2+, Zn2+, Cd2+, Zr4+) on the isotherms. The transition metal ions also showed
an increase in mean molecular area vs. DMPC on a buffered subphase without metal ions added.
In contrast to the alkaline earth metals, the transition metals show a clear difference between the
metal ion identity and the mean molecular area per DMPC monolayer. A noticeable trend in mean

110

molecular area, similar to that seen for a fatty acid monolayer, is observed: Buffer < Ni2+ ≤ Zn2+ <
Cd2+ < Zr4+.29

Figure 5-3: Surface pressure / mean molecular area-isotherm of the DMPC/NBD-LPC
monolayers formed on subphases containing buffer (red solid), and 1 mM NiCl2 (blue
dash), ZnCl2 (green dot), CdCl2 (purple dash-dot), or ZrOCl2 (orange dash). Inset:
Expanded region showing mean molecular area at dipping pressure.
We correlate the increase in mean molecular area with increased competition for the
anionic lipid phosphate moieties at the interface between the cationic metal ions in the subphase
and the pendent (cationic) choline groups. At pH = 7.5, the phosphocholine head-group will be
deprotonated leading to a net neutral30 or, as suggested by recent molecular dynamics simulations,
a slightly negative (-0.53e) charge.31-32 Regardless of the overall charge of the monolayer without
metal ions, the introduction of the positively charged metal cations (+2e) into the subphase disrupts
any preexisting electrostatically induced local organization between the pendent lipid choline

111

(+0.75e) and phosphate moieties (-1.28e). The magnitude of disruption is likely dependent upon
the strength of interaction between the metal ion and phosphonate group.33-35

Figure 5-4: Interactions between the metal ions and lipid monolayer are shown as the
mean lipid area vs. Metal ion van der Waal radius. Error bars are calculated by
extrapolation of a line tangent to the isotherm between 20-30 mN/m down to 0 mN/m.
The electrostatic competition between metal ions, anionic phosphonate, and cationic
choline are correlated as a change in the mean molecular area of the lipid monolayer for a given
metal ion’s van der Waal radius (Fig. 5-4).36 We assert that as the attractive interaction between
(cationic) metal ion and (anionic) phosphonate group increases, a concurrent increase in repulsive
interactions between either individual metal ions at the interface or metal ions and the pendant
phosphocholine should occur (dependent upon metal ion-lipid stoichiometry) resulting in the
observed expansion of the isotherms. In both instances, the induced disruption of mean molecular
area is greater for the transition metals than for the alkaline earth metals. The increased disruption

112

from the transition metal ions is likely due to the increased thermodynamic stability for the metalphosphonate transition metal complexes over their alkaline earth metal counterparts.

Electrostatic interactions mediate transfer of metal ions from subphase to support surface.
XPS was utilized to better understand the interactions mediating transfer of the metal ions to the
support surface. An idealized diagram of the supported film surface is shown in Fig. 5-5. The
values listed in Table 5-1 are relative intensities (corrected by the appropriate sensitivity factor)
for the alkaline earth and transition metals transferred to either a phosphorylated fluorine-doped
tin oxide (FTO) or phosphorylated silica surface versus phosphorus (P2p), tin (Sn3d5/2), and silicon
(Si2p) peaks.
Table 5-1: Relative metal ion surface coverage for the phosphorylated FTO and glass
surfaces are reported versus phosphorus (P2p), tin (Sn3d5/2), and silica (Si2p) peak
areas.
Glassa
FTO
Metal ion
Mx+:Sn 3d5/2
Mx+:P 2p
Mx+:Si 2p
Mg2+ (2s)c
2+
Ca (2p)
0.03
0.22
0.03
Ba2+ (3d5/2)
0.06
1.18
0.04
2+
Ni (2p3/2)
0.06
0.40
0.03
Zn2+ (2p3/2)
0.22
1.59
0.22
Cd2+ (3d5/2)
0.04
0.61
0.07
4+
b
Zr (3d)
0.80
1.07
a,b
P2p peaks were unresolved
c
Mg2s peak was unresolved. The Mg KLL auger peak (305.5 eV) is present in the Mg survey
scan, indicating transfer (Fig. A5-1)
XPS provides a semi-quantitative assessment of the presence of metal ions on the surface after
transfer from the LB-trough. For all metal ions, except magnesium, signal intensities were present
above baseline indicating that metal ions are transferred from the LB-trough to the supporting
surface (example survey scans and detailed scans are shown in Figs. A5-1, A5-2, A5-3). For a
process where the metal ion transfer to supporting surface is mediated by lipid-phosphonate

113

interactions (1:1 Mx+:DMPC) an expected ratio of Mx+:P is ca. 0.18 Âą 0.04, whereas metal ion
transfer mediated by surface phosphates (1:1 Mx+:surface phosphate) an expected ratio of Mx+:P
is ca. 0.82 Âą 0.04 (calculations can be found in appendix A5.1). The observed ratios between the
metal ion and phosphorus peaks suggests that the surface phosphates, not lipids, are primarily
mediating metal ion transfer.

Figure 5-5: Idealized diagram of the prepared supported lipid layer. The films are
comprised of the phosphorylated glass support (a.), metal ion and aqueous under-layer
(b.), and DMPC/NBD-LPC film (c.) with buffer adlayer (d.).
We note that both Zn2+ and Zr4+ have much higher ratios relative to the other metal ions.
Given the buffered pH of 7.5 and relatively low hydroxide Ksp of zinc, the increased Zn2+ ratio is
attributed to the transfer of neutral Zn(OH)2 species into the adventitious water layer described
earlier. The higher Zr4+ ratio is attributed to both complete surface phosphate coverage and also

114

due to the transfer of other soluble species such as ZrO(OH)y2-y (including nonstoichiometric
water) or hydrous oxide polymers from the trough to surface.37-38
Phospholipid Langmuir film dynamics minimally affected by metal ion identity.
Fluorescence recovery after photobleaching (FRAP) experiments were performed on the film
surfaces to better understand the influence of the metal ion identity in the under-layer on the
dynamics of the supported DMPC film. The translational diffusion of the fluorescent probe NBDLPC in the DMPC film was measured to evaluate this relationship.

Figure 5-6: Normalized FRAP recovery curve from one SiOx surface fitted to the full
reaction-diffusion model. Residuals are plotted below. Inset: Average of all trials on the
SiOx (red circles) and phosphorylated surface (blue squares) are plotted with error
(standard deviation) represented as shading.
Typical normalized recovery curves and fits for the DMPC films supported on a glass
surface are shown in Fig. 5-6. The fluorescence recovery curves were fitted to a single bindingsite model described by eqn. 5-2 where f represents the free, unbound fluorescent probe, m
115

represents a binding site, and b represents a bound complex of fluorescent probe and binding site.
The rate constants for this reaction are described as kon and koff for association and dissociation,
respectively.
kon
f +mb
koff

5-2

There are many different models used for the fitting of FRAP data and we assert that the
full reaction-diffusion model is best suited under the experimental conditions relevant to this work.
The use of a pure diffusion model that does not account for interactions between the support and
the monolayer when binding is present, such as the Soumpasis model, is inappropriate.39
Similarly, caution is also warranted in using models that have no analytical dependence on the
bleaching spot size, such as the off-rate dependent recovery described by Bulinski.40 Bulinski et
al. showed that for cases when diffusion is much faster than the binding rate constant and timescale
of the measurement, the fluorescence recovery is determined by the dissociation rate constant, koff
(eqn. 5-3).40
frap ( t ) = 1 − e

− koff t

5-3

To determine the applicability of fitting our FRAP recovery curves to the reactiondominant model described by eqn. 5-3, koff is determined as a function of bleaching spot size for
DMPC/NBD-LPC on a phosphorylated surface (Fig 5-7). The reaction dominant model relies
upon the assumption that on the timescale of the experiment, diffusion is rapid and can be
neglected, however the clear dependence of koff on spot size suggests that for our experimental
conditions, the use of the reaction-dominant recovery model is not appropriate. This finding also
suggests that for this zwitterionic lipid system, the relationship between binding rates for a given
spot size occurs on a slightly faster timescale than diffusion (i.e. k*onω2/Df >1).41

116

Figure 5-7: Bleaching spot area vs. lipid dissociation rate (koff) determined by the
reaction dominant model. Error bars represent 95% confidence interval.
Using this single binding-site model, the FRAP curves were fit using the full reactiondiffusion model described by Sprague et al. (eqn. 5-4).41
*
 1 Feq

kon
frap=
(t ) L  −
⋅ [1 − 2K1 (qω ) I1 (qω ) ] ⋅  1 +

p + koff

 p p
−1

*
 p  
kon
=
q    1 +
p + koff
 Dt  
*
kon
= kon [ m ]


 
 

12

  Beq
 − 
  p + koff

 
 
 

5-4

Where L-1 indicates the inverse Laplace transform of the bracketed function, Feq and Beq are
related to k*on and koff by eqn. A5-8, K1 and I1 are the modified Bessel functions, and ω is the radius
of the bleaching spot. (A detailed explanation of the equation and mathematics behind the model
is shown in appendix section A5.2). Using the full reaction-diffusion model yields the parameters
117

kon*, koff, and Dt where kon* and koff are the rate constants for the lipid-surface binding interactions
and Dt is the diffusion constant for the fluorescent probe in the film.

Figure 5-8: Translational diffusion coefficients acquired from fitting the full reaction
diffusion model for each prepared surface. Error bars represent 95% confidence
interval.
The diffusion constants for each surface are shown in Fig. 5-8 and Table A5-2. The
diffusion constant for the DMPC/NBD-LPC monolayer was observed to be the fastest for the
unmodified glass surface in the absence of metal ions. The slowed diffusion on the phosphorylated
surface is attributed to the increased ionic interactions of the pendent choline (cationic) with the
negatively charged phosphate groups. At pH = 7.5 the phosphorylated surface will be more
negatively charged than silica, yielding increased interaction between the pendent (cationic)
choline and surface.

118

Interestingly, for both the alkaline earth metals and the transition metal ions, the trend in
diffusion constants matches the trend in metal-ligand formation constants (Kf) for complexation
with a simple ligand (L), such as methyl-phosphonic acid (MPA) (eqn. 5-5, Fig. 5-9, assuming 1:1
M:L stoichiometry).
M + L  ML
[ ML]
Kf =
[ M ][ L]

5-5

For MPA, pKa2 is reported as 7.5 and pKa1 is reported as 2.3, while pKa1 for DMPC is
~1.42,30 Given that the phosphonic acid moiety of both MPA and DMPC contribute a -1 charge at
pH = 7.5, the interactions are expected to be similar. The same trends were also observed when
comparing the diffusion constants to aminoethane-phosphonic acid formation constants. While a
correlation between diffusion and Kf does appear to exist, the trend exists within the observed
error, indicating that the metal ion-phosphate binding plays a minimal role in driving the diffusion
in these systems.
The statistically significant difference among the specified surfaces was determined by
one-way ANOVA. Among the different alkaline metal surfaces F(2,58) = 2.0, p = 0.14 and among
differing transition metal ion surfaces F(3,71) = 3.2, p = 0.03 (Tables A5-3-A5-6). While the
observed diffusion coefficients for the transition metal ions show a significant statistical difference
and correlate with the metal-ligand formation constants (Fig. 5-9), the percent difference of the
diffusion coefficients among the metal surfaces is relatively low (22%). We discuss below how
the dominant forces mediating the fluidity in these films are lipid-lipid van der Waals dispersion
interactions.

119

Figure 5-9: Log of formation constants for methylphosphonic acid to each metal ion vs.
translational diffusion coefficient. Error bars represent 95% confidence interval.

Ionic interactions are secondary to acyl chain dispersion interactions in mediating film
fluidity. To better understand the contributions relating to the binding events occurring at the
interface between the PC head-group and the support, we used the full reaction-diffusion model to
estimate the free energy of binding for the metal-PC system eqn. 5-6,

K eq =
∆G

*
kon

0
MP − L

koff

5-6

=
− RT ln ( K eq )

An idealized mechanism for the energies required for diffusion to occur is shown in Fig.
5-10. We find that for the free energy of binding associated with the diffusional measurements
(ΔGMP-L) is ~ -4.4 ± 0.4 kJ/mol. This interaction energy is much less than typical covalent binding

120

energies and is ~3 times less than estimated dispersive energies between individual lipid tailgroups (∆GL-L ~12-18 kJ/mol-DMPC).

43-44

The translational diffusion can be considered in the

context of having two contributions to the activation energy for diffusion, one from ionic
interactions between metal and head-group (EMP-L), the other from dispersion interactions between
lipid tail-groups (EL-L), and we assert that these factors can be considered as simply additive. We
consider that the MP-L and L-L interactions are separable and thus add linearly.

Figure 5-10: Idealized representation of the energies required for the NBD-LPC to
undergo translational diffusion in a DMPC layer. The translational diffusion is governed
by both an activation energy from the metal phosphonate-lipid interaction (EMP-L) and
activation energy from the lipid-lipid van der Waals dispersion interactions (EL-L). The
free energy for translational diffusion (ΔGDt) is therefore governed by the contributions
from both metal phosphonate-lipid (ΔGMP-L) and lipid-lipid interactions (ΔGL-L).
It is important to note that changing metal ion identity affects both ΔGMP-L and ΔGL-L due
to changes in lipid-lipid spacing and thus dispersion interactions (Fig. 5-5). Using the formation
constants from MPA, we find that the free energy of binding associated with the MP-L interactions

121

is 1.1 Âą 0.5 kJ/mol and 2.4 Âą 0.3 kJ/mol for the for the alkaline earth metals and transition metal
ions, respectively. For a two-chained 14-carbon molecule separated by a distance of 8.6Å, in a
hexagonal close packing arrangement, the attractive dispersion energy is expected to be
approximately 18 kJ/mol (calculations contained in appendix section A5.5).
Accounting for changes in distances between lipid molecules (Fig. 5-5) due to changes in
metal ion identity, and approximating for changes in dispersion energy based on changes in
intermolecular distances, we find that the free energy changes associated with changes in metal
ion identity are ~6 kJ/mol and ~7 kJ/mol for the for the alkaline earth metals and transition metal
ions, respectively. Given that the energies for dispersive forces (ΔGL-L) are ~2-3 times larger than
the energies contributed from the metal to lipid interactions (ΔGMP-L), we assert that the diffusion
is mediated primarily by interactions between the DMPC molecules.

5.4

Conclusions
Despite the zwitterionic and net-neutral nature of the phosphocholine head-groups,

interactions between metal-ions introduced into a supporting subphase and the lipids are sufficient
to alter the organization and density of the monolayer. This disruption in organization is greater
for transition metals than for the alkaline earth metals examined in this study. Contrary to the
observed influence of the metal ions on the organization of the lipid monolayers, minimal
differences in diffusion were observed. The lack of influence of metal ions in the sub-layer is
attributed to the majority of diffusion being mediated by lipid-lipid interactions and van der Waals
dispersion interactions rather than by binding interactions with the metal ions and surface moieties.
Experimental studies are currently underway examining the influence of lipid-lipid interactions to
provide a cohesive and quantitative understanding of the individual contributions at play.

122

APPENDIX

123

APPENDIX

A5.1

XPS Analysis
To determine the expected ratio between metal and phosphorus mediated by the surface or

lipid, we first estimate the total number of phosphorus present:
=
PTotal PSurface + PLipid

A5-1

The content of surface phosphorus is estimated assuming complete phosphorylation of all present
surface hydroxides (5-8 OH/nm2).45-47 The phosphorus content provided from the lipids is
estimated using the known mean molecular area from the Langmuir isotherms. At 25mN/m the
average molecular area is ca. 69 Å2/molecule (0.69 nm2/molecule), at 1 P/Lipid this represents
1.4 P/nm2. The ratio of metal to phosphorus is given by:
M x+ : P =

M x+
PTotal

A5-2

Assuming a surface mediated transfer and a 1:1 ratio of metal ion to surface phosphorus:
M x + = PSurface

A5-3

Assuming a lipid mediated transfer and a 1:1 ratio of metal ion to lipid phosphorus:
M x + = PLipid

A5-4

Given these set of assumptions, the expected Mx+:P ratio for a surface mediated transfer process is
0.78 for 5 P/nm2 and 0.85 for 8 P/nm2 yielding an estimated ratio 0.82 +/- 0.04.

124

Figure A5-1: XPS survey spectra of DMPC (red), Mg2+ (blue), and Ba2+ (green) are
shown.

Figure A5-2: (left). Fit of the XPS spectral data for a DMPC/NBD-LPC film containing
Ba2+ is shown. The Ba3d5/2 peak is fit to a Gaussian-Lorentzian curve after Shirley
background subtraction. Figure A5-3: (right). Representative convoluted fits for the
overlapping Sn4s and P2p peaks on the Ba2+/DMPC/NBD-LPC film.
125

A5.2

Analysis of FRAP curves by the Full Reaction-Diffusion Model
The FRAP curves were fit using the full reaction-diffusion model described by Sprague et.

Al.41 We briefly summarize the model system described by Sprague and relate the assumptions
and equations to our lipid film system below. The analysis starts with a simple single binding site
model eqn. A5-5:

A5-5

where F =[f] represents the concentration of free diffusing molecule (NBD-LPC), M = [m]
represents the concentration of available binding site (adsorbed metal ions), and B = [b] represents
the concentration of bound complex between NBD-LPC and the adsorbed metal.
Assuming the system has reached equilibrium prior to the bleaching event, and that the
binding sites are immobile, the recovery can be described by a set of coupled reaction-diffusion
equations eqn. A5-6:

∂f
*
f + koff b
= Dt ∇2 f − kon
∂t
∂b
*
f + koff b
= kon
∂t

A5-6

where Dt is the diffusion constant of the NBD-LPC, k*on = k*on[m] is the pseudo first-order rate
constant derived by assuming that the M = [m] is constant, and ∇2 is the Laplacian operator. In
the case presented here, both assumptions are assumed valid. The surfaces were allowed to
equilibrate ~10 minutes prior to imaging and a period of 10 seconds was collected prior to the
bleaching event to assure equilibrium was reached. The binding sites in this case are considered
to be any binding interaction with the surface and the surface is assumed to be immobile on the
timescale of the measurement.
126

Because the system is at equilibrium at the start of the FRAP experiment both F and B are
at equilibrium:

df db
= = 0
dt dt
*
kon
Feq = koff Beq

A5-7

Assuming normalization of the FRAP curves, the final recovery of the free and bound species is
equal to:
Feq + Beq =
1

Feq
=

*
koff
kon
B
,
=
eq
*
*
kon
kon
+ koff
+ koff

A5-8

One way to analytically solve the reaction-diffusion differential equations is to take the Laplace
transform of the coupled equations. The inverse Laplace transform was performed using the
matlab function invlap.m on the analytical equation:

A5-9

where frap(t) is recovered by taking the inverse Laplace transform, Feq and Beq are related to k*on
and koff by eqn. 4, K1 and I1 are the modified bessel functions, ω is the radius of the bleaching spot
and q is described by eqn. A5-10:

q
=

A5.3

 p

 Df

*

kon
1
+


p + koff






A5-10

FRAP Recovery Curves and Fitting
The FRAP recoveries were fit using the matlab routine lsqcurvefit.m with Eqs. A5-9 & A5-

10 for the full reaction-diffusion model. After fitting, the FRAP recoveries were averaged for each

127

surface and are shown in Table A5-1 for the dissociation constant koff, and Table A5-2 for the
diffusion coefficient DT.
Table A5-1: Dissociation constant (koff) data acquired by fitting FRAP curves with the
reaction-dominant model.
95% Confidence
Interval for Mean
Mean
koff
Std.
Std.
Upper
Lower
Surface
N
(10-3 s-1) Dev.
Error Bound
Bound
Minimum Maximum
19
26.8
13.2
5.9
20.8
32.7
3.3
57.0
SiOx
27.2
18.3
8.0
19.1
35.2
0.1
55.4
POx-SiOx 20
19
34.3
22.3
10.0
24.2
44.3
1.0
68.6
Mg
22
32.9
20.2
8.4
24.5
41.3
0.0
75.3
Ca
18
26.7
18.7
8.6
18.0
35.3
2.5
77.0
Ba
21
39.1
20.6
8.8
30.2
47.9
2.4
75.7
Ni
25
28.9
15.4
6.0
22.8
34.9
2.8
78.0
Zn
17
29.9
24.9
11.9
18.0
41.7
1.5
84.4
Cd
9
27.1
15.7
10.2
16.9
37.3
7.7
52.1
Zr
Table A5-2: Diffusion coefficient data acquired by fitting FRAP curves with the full
reaction-diffusion model.
95% Confidence
Interval for Mean
Mean Std.
Std.
Upper
Lower
Surface
N
(DT)
Dev.
Error Bound
Bound
Minimum Maximum
19
4.50
0.72
0.32
4.18
4.82
3.53
6.17
SiOx
3.01
1.02
0.45
2.56
3.46
1.69
5.20
POx-SiOx 20
19
3.14
0.64
0.29
2.85
3.42
2.24
4.40
Mg
22
2.79
0.67
0.28
2.51
3.07
1.67
4.03
Ca
18
2.76
0.60
0.28
2.48
3.04
1.86
3.91
Ba
21
2.63
0.43
0.18
2.44
2.81
1.89
3.62
Ni
25
2.79
0.50
0.20
2.60
2.99
1.79
3.74
Zn
17
2.96
0.48
0.23
2.73
3.19
2.33
4.07
Cd
9
3.25
0.93
0.61
2.65
3.86
1.90
4.58
Zr

A5.4

One-way ANOVA Testing of the Fitted Results
One-way ANOVA analysis was performed on the series of FRAP recoveries for each

surface. The average results for each surface are shown as a box plot in Figure A5-4. The central

128

red mark represents the median and the edges of the box represent the 25th and 75th percentiles of
the data. Whiskers are shown to extend to the most extreme data points, where outliers are
represented individually. The notches in the plot correspond to the 95% confidence interval, where
if the notches of one surface do not overlap another surface, the two medians are significantly
different to 95% confidence.

Figure A5-4: Box plot for the fitted diffusion coefficients of all trials on each surface.
The red line represents median, whiskers represent maximum and minimum values, and
red asterisks are considered outliers. Notches in the box plot represent 95% confidence
intervals for each surface.
Using a Chi-Square goodness-of-fit test to the 5% significance level, trials for each surface
were normally distributed and ANOVA analysis was performed. ANOVA analyses are shown for
all surfaces (Table A5-3), among all metal surfaces (Table A5-4), among the alkaline metals only
(Table A5-5), and among the heavy metals only (Table A5-6).

129

Table A5-3: One-way ANOVA test among all surfaces.
Sum of Squares df
Mean Square
49.0
8.0
6.1
Between Surfaces
72.9
161.0
0.5
Within Surfaces
121.9
169.0
Total

Table A5-4: One-way ANOVA test among all metal surfaces.
Sum of Squares df
Mean Square
4.6
6.0
0.8
Between Surfaces
43.3
124.0
0.3
Within Surfaces
47.9
130.0
Total

F
13.5

Significance
6.9E-15

F
2.2

Significance
5.0E-02

Table A5-5: One-way ANOVA test among alkaline metal surfaces
Sum of Squares df
Mean Square
F
1.7
2.0
0.8
2.0
Between Surfaces
23.0
56.0
0.4
Within Surfaces
24.7
58.0
Total

Table A5-6: One-way ANOVA test among heavy metal surfaces.
Sum of Squares df
Mean Square
2.82
3.00
0.94
Between Surfaces
20.34
68.00
0.30
Within Surfaces
23.16
71.00
Total
A5.5

F
3.2

Significance
0.14

Significance
0.03

Interaction Energy Calculations
The interaction energy between the DMPC molecules was estimated by a summation of

the dispersive energies between two parallel (two-chained) 14-carbon alkyl chains.43-44 The
dispersion attraction energy (WDMPC) between two parallel saturated DMPC chains is given by:
WDMPC = Îą

3π N
8Îť D 5

A5-11

130

where Îą is the interaction energy between two methylene units, N is the number of methylene units
(14 for DMPC), λ is the length of the individual methylene units (1.26Å), and D is the separation
distance between chains (calculated from the mean molecular area isotherms at the transfer
pressure of 25mN/m).
The total interaction energy in the monolayer case assumes a hexagonal close packing
arrangement and is a summation of attractive energies from the first closest interacting group at
radius, r1, to the nth group at a radius of rn, where r is the interchain separation distance calculated
from the radius of the mean molecular area isotherms at the transfer pressure.
rn = 2nr
 n

WM =
2 ⋅  ∑ N nWDMPC ( rn ) − WDMPC ( r1 ) 
 1


A5-12

where Nn is the number of nearest neighbors (i.e. N1 = 6), and WDMPC(rn) is the DMPC interaction
energy at a separation distance of rn and all summations have been made to n=12.

131

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132

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136

CHAPTER 6: CONCLUSIONS AND FUTURE DIRECTIONS

137

6.1

Conclusions and Future Directions
The goal of this research has been to better understand how the molecular scale interactions

at play in a supported model membrane system affect the dynamical properties. The motivation
for this work is the creation of fluid and robust thin films with a wide range of properties for
applications such as chemical sensing, separations, self-healing membranes, molecular electronics,
and tribology. The use of these films for such applications is attractive, but is currently limited
due to the incomplete understanding of the interplay between interfacial and intermolecular
interactions that ultimately give rise to the dynamic properties. Bulk-scale properties are often
inferred from molecular scale viscosity measurements (fluorescence anisotropy decay) or bulkscale measurements are made inferring molecular scale properties. Understanding the interactions
that are operative in mediating the dynamical properties over a large range of length scales will
allow for the tailoring of these films to the desired set of applications.
The focus of the work contained within this dissertation has been on biomimetic model
membrane systems for applications in chemical sensing and separations. These systems often rely
on transmembrane proteins as the chemically selective or transducing agent. There are a variety
of methods used to prepare and examine biomimetic membrane systems including unsupported
systems such as black lipid membranes,1-3 lipid vesicles,4-7 and supported systems such as solid
supported,8-10 and hybrid bilayer membranes.11-13 This work has focused on the use of solid
supported model membranes due to the diversity of chemical means available to control the
dynamical properties such as: pH, presence and identity of metal ions in the subphase and buffer
overlayer, surface composition and roughness, as well as the necessity of a solid support for
transfer of the chemical response information of interest from the chemically selective element to
the detector.

138

In Chapter 2 of this dissertation, the instrumental methods used to examine the morphology
and dynamical properties of supported model membranes systems is described with a focus on the
measurement of rotational and translational diffusion behavior of fluorescent probe molecules.
Brewster angle microscopy (BAM) is introduced as a means to observe changes in the morphology
of amphiphilic molecules such as alkanoic and alkyl phosphonic acids, or lipids, at an air/water
interface as a function of pressure applied to form a monolayer and under a variety of chemical
and physical conditions. Fluorescence Anisotropy Decay Imaging (FADI) is introduced as a
means to spatially resolve variations in the fluorescence anisotropy decay dynamics of monolayer
or bilayer films. The information available from these measurements allow for a quantitative
measurement of the rotational diffusion properties of a fluorescent probe molecule, as well as
changes in film morphology due to compositional changes of the supporting surface or organic
monolayer.

Lastly, methodology for the examination of Fluorescence Recovery After

Photobleaching (FRAP) measurements is described. FRAP dynamics depend on the type of
fluorescent probe used, the morphology of the interfacial layer and any surface-adlayer
interactions. A variety of functional models are considered to best model the observed data, with
a goal of obtaining translational diffusion constants, and association/dissociation rate constants for
interfacial binding site events.
In Chapter 3 we reported on changes to the rotational diffusion dynamics of perylene within
a tethered lipid monolayer.14 The data revealed slowed dynamics in the lipid monolayer exposed
to air compared to more rapid diffusional dynamics when immersed in aqueous solution. The
central question in these findings is why the addition of an aqueous adlayer increases the motional
freedom of the chromophore. The decrease in rotational diffusion time constant of perylene
imbedded in a lipid monolayer covered by an aqueous overlayer is likely associated with a change

139

in the extent of organization of the lipid acyl chain region upon exposure to the aqueous overlayer.
Electrochemical data indicating ca. 52% coverage of the lipid monolayer suggests there is ample
spatial heterogeneity available for reorganization to occur. Ellipsometric and contact angle data
are consistent with a nonpolar and spatially heterogeneous lipid (sub)monolayer interface.
Unfortunately, based on the measurements made, little information is available on the length
scale(s) of the organized regions at the interface. Future studies using chromophores with different
aspect ratios (e.g. DPH)15 may shed light on the degree of lipid acyl chain conformational
organization. Additionally, at the time these measurements were made, both the FRAP and FADI
instruments were not yet available in the Blanchard lab. Reexamination of these films using
confocal fluorescence imaging (FRAP and FADI) could provide substantially more insight into
the characteristic organizational domain sizes of such interfaces.
The work described in Chapter 4 investigated the dynamics of a tethered BODIPY
chromophore contained within an alkylphosphonic acid LB film.16 The goal of this work was to
quantitate both the fluidity and adhesion strength of the organic adlayer to the interface using the
combination of translational and rotational diffusion data. By examining the diffusional behavior
of this tethered chromophore over length scales ranging from molecular (FADI measurements)
through macroscopic (Îźm-to-mm, FRAP measurements), we demonstrated a direct and novel
means for evaluating organic adlayer binding strength to a supporting surface. Using steady state
and fluorescence lifetime imaging data, we were able to determine the motional freedom that is
characteristic of the BODIPY chromophore and the effect of aggregation on molecular motion
within the ODPA film.

The data provided insight into the binding strength of a barium

phosphonate species to a silica surface and, by extension, provide an estimate of the binding
strength for related interfaces. We observed binding strengths from ∆Gdis, of ca. 20 kJ/mol. This

140

value is entirely consistent with interactions dominated by H-bonding17 and there is also an
observed chromophore concentration dependence suggesting some information on the interaction
energy of the aggregated chromophores.
The strength of interaction for our monolayer containing barium with a silica support is
very similar to the binding interactions responsible for the formation of alkanethiol monolayers on
gold and mercury.18-20

This finding is important because the spectroscopic evaluation of

chromophore dynamics in an alkanethiol/gold SAM are limited by chromophore quenching due to
the presence of a metal support,21 and this method offers a way to estimate the expected dynamics
for this family of systems.
Studies examining the role of chemical modification of the support as a means of
controlling binding strength were performed in Chapter 5, however there remain a few unanswered
questions pertaining to the system at hand. Even at the lower chromophore concentrations, some
aggregation was present. The aggregation of chromophore is likely due to the dissimilar chemical
structure between the ODPA and BODIPY probe molecule. To mitigate aggregation, preparation
of a film with a chromophore similar to the chemical structure of the film is suggested for future
studies. In addition, the combination of barium and ODPA was found to provide measurable
diffusional properties on the timescale of minutes. Examination of a combination of ODPA and
other metal systems were examined including Ca2+ and Cd2+, however diffusion was slow (tens of
minutes) and very little recovery was observed. These observations provide some key guidelines
in the preparation of fluid and robust films, which will be discussed below.
Chapter 5 examined the role of chemical modification of the support via combination of
phosphorylation of the support surface and varying the metal ion identity between the
phosphorylated surface and organic adlayer as a means of controlling binding strength.22 X-ray

141

photoelectron spectroscopy (XPS) was utilized to measure semi-quantitatively the amount of metal
ions transferred during deposition of the films. Fluorescence recovery after photobleaching
(FRAP) revealed the translational diffusion properties of the BLPA chromophore after deposition
onto a solid support. The addition of metal ions into the subphase was sufficient to alter the
organization and density of the amphiphile monolayer at the air/water interface, despite the netneutral character of the zwitterionic DMPC lipid. The disruption was found to be significantly
greater for the transition metal ions relative to the alkaline earth metals examined in the study.
Despite the observed influence of the metal ions on the organization of the lipid monolayers at the
air/water interface, minimal differences in diffusion were observed upon transfer to a supporting
surface. Given the observed transfer to, and presence of the metal ions at the supporting interface,
the lack of influence of metal ions is attributed to diffusion being mediated by lipid-lipid
interactions and van der Waals dispersion interactions.
The work presented in this dissertation shows both the ability to measure quantitatively the
fluidity and binding strengths associated with supporting model membrane systems at a solid
interface, and hints toward the ability to control the fluid and adhesion properties based on
modification of the solid support.

This work provides a few key lessons to guide future

investigations of these types of systems. (i.) The addition of an aqueous adlayer at the organic
interface disrupts organization of the alkyl region, but is essential for diffusional measurements to
be made. (ii.) Chromophores should be chosen with chemical structure similar to the amphiphile
used in the film formation if aggregation is to be mitigated. (iii.) The addition of different metal
ions into the subphase shows promise as a means of modulating film fluidity and adhesion.
One area for future investigation should be to understand the role of the lipid-lipid chain
interactions relative to the binding constants to the metal ions. This can be accomplished by

142

changing the charge associated with the amphiphile headgroup, changing the lipid chain length, or
changing the degree of saturation in the lipid acyl chain region to increase interchain distances and
thereby limit dispersion interactions.

Additional studies can be performed examining the

temperature dependence of dynamics and morphology in these systems. Changes in temperature
will likely give rise to changes in surface-lipid, lipid-lipid, and lipid-chromophore interactions.
Further changes in the supporting surface charge or spatial charge distribution via changes in the
metal oxide used for the support, or electrical bias of the interface could provide further control
over the fluidity and adhesion. A quantitative understanding of the interplay between the physical
interactions involved at these interfaces will open the door to a general way to tailor systems with
desired adhesion and diffusion properties for the required application.

143

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144

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