ABSTRACT 
 
Title of Document: Blueprinting Self-Assembled Soft Matter: An ?Easy? 
Approach to Advanced Material Synthesis in Drug 
Delivery and Wound Healing 
    Matthew Dowling, Doctor of Philosophy, 2010 
 
Directed By:    Professor Srinivasa R. Raghavan, Department of Chemical  
    and Biomolecular Engineering 
 
 
From Jello to mayonnaise to silly putty to biological cells, our world is replete with ?soft 
matter? ? materials that behave as soft, deformable solids or highly viscoelastic liquids. 
Living systems, in particular, can be thought of as extremely sophisticated ?soft? 
machines, with each cellular unit representing a touchstone for the functional potential of 
soft materials built via self-assembly. Drawing inspiration from biology, we blueprint 
soft biomaterial designs which rely upon self-assembly to achieve enhanced functionality. 
As opposed to complex synthesis schemes often used to develop improved biomaterials, 
we take an ?easy? approach by allowing relatively simple molecules orchestrate 
themselves into advanced machines. In this dissertation, we describe four separate ?soft? 
systems, all constructed by self-assembly of amphiphilic molecules under designed 
and/or triggered conditions in aqueous media. These systems revolve around a common 
theme: the structural tandem of (1) vesicles and (2) biopolymers, and the resulting 
interactions between the two. Our blueprints show promise in several important 
biomedical applications including controlled drug release, tissue engineering, and wound 
care. 
 
In the first part of this study, we blueprint a biopolymer gel that entraps pH-sensitive 
vesicles. The vesicles are formed by the self-assembly of a single-tailed fatty acid 
surfactant. We show that the gel has pH-responsive properties imparted upon it via the 
embedded vesicle nanostructures. Specifically, when the gel is brought in contact with a 
high pH buffer, the diffusion of buffer into the gel disrupts the vesicles and transforms 
them into micelles. Accordingly, a vesicle-micelle front moves through the gel, and this 
can be visually seen by a difference in color. The disruption of vesicles means that their 
encapsulated solutes are released into the bulk gel, and in turn these solutes can rapidly 
diffuse out of the gel. Thus, we can use pH to tune the release rate of model drug 
molecules from these vesicle-loaded gels into the external solution.  
 
In the second part, we have blueprinted hybrid biopolymer capsules containing drug-
 loaded vesicles by means of a one-step self-assembly process. These capsules are called 
?motherships? as each unit features a larger container, the polymer capsule, carrying a 
payload of smaller vesicular containers, or ?babyships,? within its lumen. These 
motherships are self-assembled via electrostatic interactions between oppositely charged 
polymers/surfactants at the interface of the droplet. Capsule size is simply dictated by 
drop size, and capsules of sizes 200-5000 ?m are produced here. Lipid vesicles, i.e. the 
babyships, are retained inside motherships due to the diffusional barrier created by the 
capsule shell. The added transport barrier provided by the vesicle bilayer in addition to 
the capsule shell provides sustained drug release from the motherships. Furthermore, this 
one-step drop method allows for the rapid synthesis of soft materials exhibiting structural 
features over a hierarchy of length scales, from nano-, to micro- to macro-.  
Thirdly, we have therapeutically functionalized biopolymer films by simply passing a 
solution of vesicles over the film surface. We deposit films of an associating biopolymer 
onto patterned solid substrates. Subsequently, these polymer films are able to 
spontaneously capture therapeutically-loaded vesicles from solution; this is demonstrated 
both for surfactant as well as lipid vesicles (liposomes). Importantly, it is verified that the 
vesicles are intact ? this is shown both by direct visualization of captured vesicles (via 
optical and cryo-transmission electron microscopy) as well as through the capture and 
subsequent disruption of drug-filled vesicles. Such therapeutically-functionalized films 
may be of use in the treatment of chronic wounds and burns. 
  
Lastly, we have demonstrated that the addition of a certain biopolymer transforms a 
suspension of whole blood into a gel. This blueprint is inspired from previous research in 
our group on the biopolymer-induced gelation of vesicles, which are structurally similar 
to cells. Upon mixture with heparinized human whole blood, this amphilic biopolymer 
rapidly forms into an ?artificial clot.? These mixtures have highly elastic character, with 
the mixtures able to hold their own weight upon vial inversion. Moreover, the biopolymer 
shows significant hemorrhage-controlling efficacy in animal injury models. Such 
biopolymer-cell gelation processes are shown to be reversed via introduction of an 
amphiphilic supramolecule, thus introducing the novel concept of the ?revesible 
hemostat.? Such a hemostatic functionality may be of large and unprecedented use in 
clinical the treatment of problematic hemorrhage both in trauma and routine surgeries. 
 
 
 
BLUEPRINTING SELF-ASSEMBLED SOFT MATTER:  
AN ?EASY? APPROACH TO ADVANCED MATERIALS SYHNTHESIS IN DRUG 
DELIVERY AND WOUND HEALING 
 
 
Matthew B. Dowling 
 
 
 
 
Dissertation Submitted to the Faculty of the Graduate School of the 
University of Maryland, College Park, in partial fulfillment 
of the requirements for the degree of 
Doctor of Philosophy 
2010 
 
 
 
 
 
 
 
 
 
Prof. Srinivasa R. Raghavan, Dept. of Chemical and Biomolecular Engineering, Chair 
Advisory Committee: 
Prof. William E. Bentley, Fischell Dept. of Bioengineering 
Prof. Philip R. DeShong, Dept. of Chemistry and Biochemistry 
Prof. John P. Fisher, Fischell Dept. of Bioengineering 
Prof. Gregory F. Payne, Dept. of Bioengineering 
 
 
 
 
 
 
 
 
 
 
 
 
 
? Copyright by 
Matthew Dowling 
2010 
 
 ii 
Acknowledgements 
First and foremost, I would like to thank my advisor Dr. Srinivasa Raghavan. I 
simply cannot begin to thank him for the past five years. He has not only been the most 
influential person on my scientific career, but on my entire outlook in life. His influence 
has been massively positive on my graduate career and has provided an amazing 
foundation from which to advance my career. Being able to work in his group has been 
an unbelievable blessing. His inspiration, outlook and enthusiasm are infectious, and it 
indeed translates into the impressive production and attitude of his group members. I owe 
all of my successes in grad school to him. He is truly a model educator. 
Next, I would like to thank my family for their unending support. I am 
particularly grateful to my parents, Tom and Sandy, who have instilled in me a priority of 
extremely high value on education. Only in graduate school have I been able to truly see 
why education really is the foundation for the future, and I thank them for force-feeding 
that to me early in life. Also, I thank my siblings, Laura, Chris, Tommy, Stephen and 
William, for their belief in me in whatever I pursue, and for their humor when I need a 
laugh. I believe in all them and I hope that I can inspire them in the way that they inspire 
me.  
I would also like to thank my committee: Dr. William Bentley, Dr. John Fisher, 
Dr. Philip DeShong, and Dr. Greg Payne. I?ve collaborated with every one their labs at 
some point or another in the past five years. All of them have done world-class research 
that I could not help but be interested in learning more about during my time here at 
UMD. They have all developed some amazing grad students who share their advisors? 
 iii 
love of learning and collaborative spirit. This is a testament to what makes Clark School 
such a great place to do graduate research and why the Fischell Department, in particular, 
is moving up the national ranks so quickly. Thank you to my collaborators at University 
of Maryland, Baltimore, Dr. Hamid Ghandehari (now at Univ of Utah) and Dr. Anjan 
Nan in the Pharmacy School, and Dr. John Hess and Dr. Grant Bochicchio at the School 
of Medicine. Working with these researchers has allowed me to gain absolutely 
invaluable clinical insight into the problem that I frame from a basic materials 
development standpoint.  
I am infinitely grateful to all of my past and present fellow labmates in Dr. 
Raghavan?s lab: Jaeho Lee, Hee-Young Lee, Shihuang Tung, Bani Cipriano, Aimee 
Ketner, Rakesh Kumar, Kunshan Sun, Chao Zhu, Oluwatosin Ogunsola, Peter Thomas, 
George Chacko, Khyati Tiwari, Vishal Javvaji, Veidhes Basrur, Kunqiang Jiang, Neville 
Fernades and Charles Kuo. Without you guys, my research would amount to nothing. 
This is not to mention some outstanding undergrads I have had the pleasure of working 
with as a grad mentor including Mark Keibler, Patrick Kelleher, Nick Levy, Leigh Quang, 
Shelby Skoog, Datren Chiou, Gabrielle Galvez, Joe Bender, Jerry Wierwille and Alex 
Gedra. A number of them have gone on to top 10 graduate engineering programs, and 
that makes me truly proud.  
I certainly have to give a great deal of special thanks to my Remedium 
Technologies co-founders, Peter Thomas, Oluwatosin Ogunsola, Bani Cipriano and Chao 
Zhu. In some sense, I feel like we have achieved the ?impossible? by starting a biotech 
company as grad researchers. The undying enthusiasm of these colleagues has helped 
push me to achieve feats of which I thought I was not capable; not the least of which are 
 iv 
a number of instances of getting back ?in the saddle? after some very frustrating failures. 
Furthermore, MTECH has been massively instrumental in supporting Remedium via the 
MIPS and VA programs. Special thanks to Martha Connelly, Jim Chung, Dean Chang, 
Akbar Dawood, John Shin, Herb Rabin and Karen Thornton. Without the support and 
guidance of these individuals, the company would simply be non-existent.  
 I have to acknowledge my great friends and fellow musicians/entrepreneurs, 
Daniel Scheuerman, Brian Hospital, Dominic Campanaro and Chris Scheffey. I?ve 
learned so much from these guys and been through so many unforgettable experiences. 
Your insight into life is consistently inspiring. Your humor is addictive. Your talents are 
endless.  
 I deeply thank my girlfriend, Katie Horner, who has been my rock over the past 
many years. Without her support, I?d be nowhere. I?m excited for our future together and 
for her upcoming graduate career at NYU. 
Last, but not least, I thank Dr. Robert Fischell for his support of my graduate 
research and entrepreneurship via the Fischell Fellowship in Biomedical Engineering. His 
inspiration and example has driven me over the years to work relentlessly in both science 
and business, and to learn how the two can work together to create real solutions for the 
improvement of human health. I?m forever indebted to his generosity and his spirit.  
 
 
 
 
 v 
TABLE OF CONTENTS 
 
Acknowledgements ??????????????????????????  ii 
 
Table of Contents ?????????????????????????.......  v 
 
List of Tables .........?................................................................?.......................??.. vii 
 
List of Figures .........?................................................................?.......................??viii 
 
1. Introduction and Overview ....................................................................???......... 1 
1.1.  Problem Description and Motivation.??????????????.............. 1 
1.2.  Our Approach .............................................???????????.................3 
1.3.   Significance of this Work???????????????????...8 
 
2. Background ........................................................???????..........???........11 
2.1.  Vesicles and Liposomes ?...........??......????????........................... 11 
2.2.  Associating Polymers................................?????????..........?..17 
2.3. Biopolymers ??????????????????????????.. 19 
2.4 Cyclodextrins ???????????????????????............ 24 
2.5.  Characterization Technique ? I: Rheology....................................??????.26 
2.6. Characterization Technique ? II: SANS ????????????????30 
2.7.   Characterization Technique ? III: DLS??????????????.32 
2.8.   Characterization Technique ? IV: Cryo-TEM???????????..33 
 
3. Biopolymer Gels Containing pH-Sensitive Vesicles  ..........????..???..? 36 
3.1.  Introduction ........?...................???....................................................... ?? 36 
3.2.  Experimental Section .........??........................................................................... 40 
3.3.  Results and Discussion ...........................................??.?................................. 42 
3.4.  Conclusions ....................................???..?.......???????................. 57 
 vi 
 
4. Biopolymer Capsules Containing Vesicles  ..??. ?????...............................58 
4.1.  Introduction ........?...................???....................................................... ?? 58 
4.2.  Experimental Section .........??........................................................................... 65 
4.3.  Results and Discussion ...........................................??.?......................... ........70 
4.4.  Conclusions ....................................???..?.......???????......... ........ 80 
 
5. Vesicles Capture on Patterned Surfaces Coated with Amphiphilic Biopolymer.. 82 
5.1.  Introduction ........?...................???....................................................... ........ 82 
5.2.  Experimental Section .........??............................................................................86 
5.3.  Results and Discussion ...........................................??.?..................................90 
5.4.  Conclusions ....................................???..?.......???????................ 102 
 
6. Gelling Blood with an Amphiphilic Biopolymer.....................................................104 
6.1.  Introduction ........?...................???....................................................... ...... 104 
6.2.  Experimental Section .........??................................................................... .... 108 
6.3.  Results and Discussion ...........................................??.?........................ ......112 
6.4.  Conclusions ....................................???..?.......???????......... ......122 
 
 
7. Conclusions and Recommendations............?.??????..........?????. 124 
7.1.  Conclusions ???????????..???????............................... 124 
7.2. Recommendations for Future Work ???????????????........126 
 
8. References .....................................................?.??????????????130 
 
 
 
 
  
 vii 
LIST OF TABLES 
Table 2.1. Properties of the three naturally occurring cyclodextrins                         26 
 
Table 3.1. Parameters obtained by fitting Peppas Equation to dye release curves    56 
 viii 
LIST OF FIGURES 
Figure 1.1. Overview of the Four Blueprinted Self-Assembled Soft Systems. (a) 
Controlled-Release Jello: pH-Sensitive Vesicles within Biopolymer Gels, (b) 
?Mothership? Drug Carriers: Biopolymer Capsules Containing Vesicles; (c) Band-Aids 
for Chronic Wounds: Biopolymer Films Capturing Vesicles; (d) Self-Assembled 
Hemostats: Associating Biopolymers Gelling Vesicles/Cells                                           4 
 
 
Figure 2.1.  The structure of vesicles formed by the self-assembly of amphiphiles. The 
vesicle is formed by the folding of an amphiphilic bilayer that is about 2-5 nm in 
thickness.                                                                                                                            12 
 
Figure 2.2. Schematic illustration of the formation of vesicles by self-assembly of 
amphiphilic molecules in water. Due to their cylindrical geometry, lipids, which are two-
 tailed amphiphiles, can form vesicles in aqueous solution upon folding of lipid bilayers 
into spheroid structures represented in the cartoon above. Also, pairs of fatty acid 
surfactants can collectively take on a cylindrical geometries and, thus, form vesicles in 
the same way as lipids.                                                                                                       14 
 
Figure 2.3. Preparation of unilamellar lipid vesicles of various sizes: (a) small or large 
vesicles (20 nm ? 200 nm);20 (b) giant vesicles by electroformation                                17 
 
Figure 2.4. Archiecture of a telechelic associating polymer and the structures formed by 
its self-assembly in aqueous solution                                                                                 18 
 
Figure 2.5. Structures of the parent sugars in (a) chitin and (b) chitosan. The N-acetyl-D-
 glucosamine sugar in chitin is deacetylated to give the D-glucosamine sugar in chitosan. 
                                                                                                                                            21 
 
Figure 2.6.  Structure of hydrophobically-modified chitosan (hm-chitosan) with C12 
hydrophobic tails.                                                                                                               22 
 
Figure 2.7. Structure of Gellan Gum                                                                                 24 
 
Figure 2.8. Truncated cone-shaped conformation of ?-CD                                              25 
 
Figure 2.9.  Photograph of the Rheometer RDA-III strain-controlled rheometer being 
operated in a cone-and-plate geometry.                                                                             27 
 
Figure 2.10.  Schematic of a SANS experiment.                                                              30 
 
Figure 2.11.  Photograph of the controlled environment vitrification system (CEVS) used 
for sample preparation in cryo-TEM. The schematics on the left show the various steps in 
the process.                                                                                                                        35 
 
 ix 
Figure 3.1. Photographs and titration curve for 1% NaOA at 25?C. Increasing amounts of 
1 M HCl are added to a micellar solution of NaOA (pH 10) and the corresponding 
solution pH at equilibrium is shown in the plot. Structural assignments are done in 
accordance with Ref. 10. Photographs of samples corresponding to different regions of 
the plot are shown above.                                                                                                   43 
 
Figure 3.2. Photographs and schematics of (a) gelatin gel; (b) NaOA vesicles; (c) gelatin 
gel loaded with NaOA vesicles. The gelatin gel is a 3-D network of gelatin chains, with 
chain segments connected into triple helices at the crosslink points. When vesicles of 
diameter ~ 100 nm are entrapped in the gelatin gel, the initially colorless gel assumes a 
bluish hue due to light scattering from the vesicles.                                                          45 
 
Figure 3.3.  SANS data at 25?C for 0.5% NaOA vesicles at pH 8.3 (circles) and a 5% 
gelatin gel loaded with 0.5% NaOA vesicles (squares). In both cases, the intensity I 
follows a slope of ?2 at moderate q, which is characteristic of scattering from vesicles.  
                                                                                                                                            47 
 
Figure 3.4.  Movement of micellar front within a vesicle gel due to diffusion of pH 10 
buffer. The buffer is introduced into the headspace above the gel at time zero. As the 
buffer diffuses into the gel, NaOA vesicles in the gel are converted into NaOA micelles, 
as indicated by the change in a portion of the gel from bluish to colorless. As time 
progresses, the micellar front travels deeper into the gel, indicating that more of the 
vesicles are converted into micelles. After 9 h, the micelle region covers most of the gel. 
The plot on the right shows the interface position (measured from the top of the vial) as a 
function of time. The line through the data is a fit to eq 1.                                                49 
 
Figure 3.5. Patterning of vesicle gels by inducing localized vesicle to micelle transitions. 
(Top panel): core-shell structure with a vesicle-rich core surrounded by a micelle-rich 
shell. This is accomplished by immersing a spherical gel (41 mm diameter) in a pH 10 
buffer. The photographs correspond to increasing incubation times in this buffer solution. 
(Bottom panel): gel in a petri dish with micelle-rich regions created at discrete points. 
This was done by sticking straws into the gel and subsequently filling the straws with the 
pH 10 buffer. The photographs again correspond to increasing times and show the 
expansion of the micelle-rich regions as the buffer diffuses into the gel.                         51 
 
Figure 3.6. Release profiles of calcein dye from a gelatin gel and from a vesicle-loaded 
gelatin gel. In the case of the vesicle-gel, the dye was encapsulated within the vesicles, 
which were then embedded within the gel. The data show a slower release of dye from 
the vesicle-gel relative to the control gelatin gel. After 10 h, Triton X-100 detergent was 
added to the external solution for both samples (this point is marked by the arrow). The 
detergent diffused in and disrupted the vesicles, which lead to an increase in the rate of 
dye release.                                                                                                                         53 
 
 
 
 x 
 
Figure 3.7.  Tunable calcein release from a vesicle-loaded gel based on the pH of the 
external solution. The control is a gelatin gel surrounded by a pH 8.3 buffer and in this 
case (red circles), the dye is released rapidly. The same gel loaded with vesicles releases 
dye much more gradually at pH 8.3 (green hexagons): in this case, the vesicles are intact 
and the vesicle bilayer thus presents a transport resistance. On the other hand, if the same 
vesicle-loaded gel is placed in contact with pH 10 buffer, the dye release is more rapid 
(blue triangles): in this case, the high pH converts the vesicles into micelles, thereby 
eliminating the transport resistance due to the vesicle bilayers. All the data are fit to eq 2 
and the fit parameters are shown in Table 1.                                                                     55 
 
Figure 4.1. Formation Chitosan ?mothership? capsules containing vesicles. Capsules 
are formed by dropping a positively-charged amphiphilic biopolymer mixed with vesicles 
(~100 nm in size) into a solution of negatively charged SDBS surfactant or negatively 
charged gellan gum biopolymer. Spherical capsules are formed instantaneously. Capsule 
size is dictated by droplet size and sizes can range from ~100 ?m (micro) to 10,000 ?m 
(macro)                                                                                                                               62 
 
Figure 4.2. Schematic of Chitosan ?mothership? capsules attacking a tumor. In step 
(1), the microcapsule ?motherships? would be docked at the tumor site using antibodies 
or magnetic targeting and thereafter release one drug from the capsule lumen (e.g. an 
anti-angiogenic drug). In step (2), capsule shell would become ruptured due to enzymatic 
degradation, and ?babyship? vesicles carrying cytotoxic drug would escape freely into the 
tumor  tissue.                                                                                                                      64 
 
Figure 4.3. Controlled Release of Dye from ?Mothership? Capsules. In (a), a sustained 
release is observed for chitosan-gellan capsules impregnated with carboxyfluorescein-
 loaded liposomes (red circles). Chitosan-gellan control capsules with dye freely dispersed 
inside the capsule lumen shows a ?dosage dumping? release profile (blue circles). 
Addition of Triton X-100 at 3.25 h shows a spike in dye release for ?mothership? 
capsules, indicating vesicle intactness. In (b), chitosan-SDBS capsules impregnated with 
CF-loaded liposomes show (blue circles) show a similar release profile to control 
capsules containing no liposomes (yellow triangles), suggesting potential liposome 
rupture during capsule formation. In contrast, hm-chitosan capsules formed in SDBS 
while retaining liposomes, shows a sustained release over a 2 hour interval.                   72 
 
Figure 4.4. Magnetically targeted ?mothership? capsules via encapsulated ferrite 
nanoparticles show their magnetic properties in response to a bar magnet. (a) magnet 
placed near capsules in a vial (b) magnet placed next to a tube in which the capsules are 
flowing along with the fluid (water) from left to right.                                                     74 
 
Figure 4.5. Antibody targeted ?Motherships. (a) Fluorescently tagged primary IgG 
antibody (emission 600 nm) is covalently conjugated to the surface of a Chitosan-SDBS 
?mothership? capsule as shown by fluorescence micrograph. After incubation with a 
secondary anti-IgG antibody and subsequent rinsing, fluorescent signal at 520  nm is 
 xi 
observed, indicating that a significant amount of primary surface IgG is actively available 
for ?lock-and-key?binding.                                                                                               75 
 
Figure 4.6. Enzymatic Degradation of ?Mothership? Capsules. (a) Phase contrast 
micrograph of Chitosan ? Gellan capsules formed by standard drop method via 22G 
syringe needle; (b) 1h, 2d, and 5d micrographs of chitosan-gellan capsule after addition 
of 10 units/ml of chitosanase in external buffer solution. Capsule appears almost wholly 
degraded after the studied 5 day interval. Note that the chitosanase cannot degrade 
chitosan capsules formed with SDBS.                                                                               77 
 
Figure 4.7. Schematic of hierachical nano-micro-macro ?motherships.? By a simple 
one-step self-assembly process, structures displaying a hierchary of length scales, from 
nano (i.e. vesicles), to micro (microcapsules), to macro (large capsules) can be produced. 
These structures can package a wide variety of molecules and targeting agents either on 
their surface, or within their multi-level lumens.                                                               78 
 
Figure 4.8. Phase Microscopy (2.5?) of Capsules-within-capsules. Small Chitosan 
SDBS capsules were formed by the standard drop method via 22G syringe needle. After 
incubation, capsules were removed from SDBS solution and placed into a fresh chitosan 
solution. Chitosan solution containing capsules was uptaken by a wide-aperture fluid 
transfer pipet and dispensed dropwise into a solution of SDBS, thus forming capsule-
 within-capsule structures.                                                                                                  79 
 
Figure 4.9. Release of dye from capsules-within-capsules. Release of 
carboxyfluorescein dye from smaller capsules packaged with larger capsules (red circles) 
is significantly sustained over a 24 period relative to the same number of small capsules 
freely dispersed in aqueous buffer (blue circles).                                                              80 
 
Figure 5.1. Schematic showing the spatiotemporal capture of vesicles using hm-
 chitosan. The structure of hm-chitosan as well as a schematic of its chain are shown in 
the top left. The chain has a hydrophilic backbone shown in blue and pendant 
hydrophobes (C12 tails) that are shown in red. In step (1), thin films of hm-chitosan are 
electrodeposited onto gold cathodes that are patterned on a silicon chip. Next, in step (2), 
the chip is incubated with a solution of vesicles. Intact vesicles become spontaneously 
anchored onto the polymer films. The inset shows the likely mechanism for such 
anchoring, which is via non-covalent interaction between the hydrophobes on the 
polymer and vesicle bilayers.                                                                                            85 
 
Figure 5.2. Photograph of micropatterned chips used in this study. The leads and 
electrodes are formed by depositing gold on a silicon wafer using lithographic techniques. 
On the left is a 2-electrode chip and on the right a 6-electrode chip. A US penny is shown 
for size comparison.                                                                                                           92  
 
 
 
 
 xii 
Figure 5.3: Comparison of hm-chitosan (H) and chitosan (C) films in terms of their 
ability to capture vesicles. Top images: Green-filtered fluorescence micrographs showing 
deposition of NHS-tagged hm-chitosan on the left electrode and NHS-tagged chitosan on 
the right electrode of a two-electrode chip. The chips are then incubated with vesicles for 
10 min, followed by rinsing with DI water. Bottom images: Red-filtered fluorescence 
micrographs showing the presence of DiI-tagged vesicles on the electrodes. The results in 
(a) (left-column) are obtained with catanionic surfactant vesicles, while those in (b) 
(right-column) are obtained with liposomes. In both cases, the vesicles are anchored 
preferentially to the hm-chitosan film than to the chitosan one.                                        94 
 
Figure 5.4: Variation in the extent of vesicle capture based on the time of chip incubation 
with a vesicle solution. Hm-chitosan (H) and chitosan (C) are deposited on the left and 
right electrodes, respectively. The chip is incubated with a solution of surfactant vesicles 
for different lengths of time and then imaged by fluorescence microscopy. The images 
are red-filtered fluorescence micrographs showing DiI-tagged vesicles on the electrodes. 
A constant exposure time of 20 s at the excitation wavelength was used to obtain each 
image. The results show increasing capture of vesicles on the H side and negligible 
capture on the C side.                                                                                                         96 
  
Figure 5.5: Immobilization of giant unilamellar vesicles (GUVs) on an hm-chitosan film. 
After 10 min incubation with DiI-tagged GUVs, multiple red-fluorescent ?hot spots? 
appear on the electrode bearing hm-chitosan. A zoomed-in image of a ?hot-spot? shows 
an intact GUV of ca. 20 ?m diameter.                                                                              97 
 
Figure 5.6: Evidence for intact capture of liposomes on hm-chitosan films. (a) Green-
 filtered fluorescence image showing preferential binding of liposomes (filled with the 
hydrophilic dye CF) to the hm-chitosan (H) side relative to the chitosan (C) side. The 
chip is then exposed to the detergent Triton X-100 and rinsed. The detergent disrupts the 
liposomes into micelles, as shown by the schematics, and the encapsulated dye is thus 
released and washed away. (b) Fluorescence image of the rinsed chip shows nearly 
complete loss of fluorescent signal, confirming disruption of originally intact liposomes.     
                                                                                                                                            98 
 
Figure 5.7: Cryo-TEM image of vesicles on an hm-chitosan film. The hm-chitosan was 
deposited on the TEM grid and then used to capture surfactant vesicles from solution. 
The image shows numerous spherical objects of ~ 100?150 nm diameter, which are 
presumably intact vesicles.                                                                                                99 
 
Figure 5.8: Patterning of vesicles on 6-electrode chips. (a) NHS-tagged hm-chitosan (H) 
and chitosan (C) are electrodeposited in an alternating pattern, HCHCHC. (b) Following 
contact with DiI-tagged surfactant vesicles and rinsing, the red fluorescence indicative of 
vesicles is found only on the H electrodes. (c) H and C are electrodeposited in an 
?outside-inside? pattern, HCHHCH. (d) Once again, DiI-tagged surfactant vesicles are 
found anchored only on the H electrodes.                                                                       100 
 xiii 
Figure 5.9: Protein sensing via vesicles anchored to an hm-chitosan film. (a) Schematic 
of the experiment: biotin-decorated liposomes are first immobilized, then the chip is 
contacted with the fluorescently labeled protein, FITC-streptavidin. The binding of the 
protein is then detected by fluorescence microscopy. (b) Fluorescence image showing a 
strong signal from the hm-chitosan (H) electrode compared to the chitosan (C) electrode. 
The strong signal is indicative of protein binding to the biotins on the anchored liposomes. 
(c) Analysis of the intensities in (b). The average gray values for each electrode area are 
listed above the corresponding regions.                                                                           102 
 
Figure 6.1. Schematic of hydrophobically-modified (hm)-chitosan as a self-
 assembling hemostat. When adding hm-chitosan to a whole blood cell suspension, the 
resulting mixture self-assembles into a 3-dimensional network. This self-assembly is 
driven by insertion of hydrophobes into blood cell membranes and the subsequent 
physical crosslinking of adjacent cells.                                                                           107 
 
Figure 6.2. Comparison of the native and hydrophobically-modified polymers with 
regard to their physical influence on blood. In (a), photographs show hm-chitosan at 
0.25 wt% mixed with heparinized human whole blood (50 wt %) is able to hold its 
weight upon test tube inversion, whereas unmodified chitosan at 0.25 wt% mixed with 
heparinized human whole blood (50 wt %) remains as a liquid. A cartoon of the proposed 
?bridging? effect of hm-chitosan on blood cells is shown above the left photograph; a 
contrasting schematic of unmodified chitosan mixed with blood is above the right 
photograph. In (b), steady shear rheology displays a dramatically increased (~O(105)) 
zero-shear viscosity of the hm-chitosan-blood mixture (red circles), relative to both hm-
 chitosan (0.25 wt%) with no blood (cyan hexagons) and chitosan-blood (yellow triangles) 
controls. In (c), dynamic rheology of the hm-chitosan blood mixture (G?, closed red 
circels, G? closed green triangles) is that of gel-like elastic material which relaxes slightly 
over time, whereas rheology of the unmodified chitosan/blood mixture (G?, open green 
triangles, G?, open red circles) is that of a viscous sol                                                    112 
 
Figure 6.3. Time-to-gelation of hm-chitosan/blood mixtures. In (a), a time-sweep of 
an hm-chitosan (0.25%)/blood (50%) mixture displays the characteristics of an elastic 
material (G? [red triangles] > G? [green triangles]) over the studied interval of 11 min. 
G? > G? immediately at t = 0 min for this sample. In (b), unmodified chitosan 
(0.25%)/blood (50%) mixture shows that the sample remains as a viscous sol (G? [green 
triangles] > G? [red triangles]) over the same interval. Lastly, in (c), a sample comparison 
in a rat femoral vein injury model demonstrates that hm-chitosan (HM-CS) solution 
(0.5%) significantly decreases bleeding time relative to saline buffer (SAL) and native 
chitosan (CS) solution  (0.5%) controls.                                                                                        
114 
 
 
 
 
 xiv 
 
Figure 6.4. Application of an hm-chitosan sponge to a porcine femoral artery 
puncture. In (a) the injury was created in the femoral artery, resulting in high-pulsatile 
flow. An hm-chitosan is then applied to the injury in (b) by direct compression. After 2 
min of compression, the bandage is observed in (c) to be attached to the wound site and 
providing hemostasis. The bandage is removed after 180 min (d) with no rebleeding 
during the studied interval.                                                                                              117 
 
Figure 6.5. Dynamic rheology of hm-polymer mixed with separated blood 
hematocrit and plasma. The hm-chitosan concentration is maintained at 0.25%. 
Hematocrit was resuspended in isotonic buffer after centrifugation; plasma was removed 
and stored for use. Visual observation of an hm-chitosan-hemotacrit mixture shows that 
the sample can hold its weight upon vial inversion, whereas an hm-chitosan-plasma 
mixture flows freely as a viscous liquid. Dynamic rheology confirms the visual evidence, 
as the hematocrit mixed with polymer (G? [closed red circles] and G?[closed green 
circles]) is that of a gel-like elastic material which relaxes slightly over long times. In 
contrast, the rheology of the plasma mixed with polymer (G? [open red triangles] and 
G?[open green triangles]) is close to that of a sol-gel transition.                                    119 
 
Figure 6.6. Reversal of blood-gelation via ?-cyclodextrin (CD). In (a), a cartoon of ?-
 CD displays its supramolecular barrel-shaped structure, with hydrophobic core and 
hydrophilic exterior. A schematic hypothesis shows that by adding these molecules 
together with hm-chitosan-blood mixtures, the samples become liquefied due to 
?capping? of hydrophobes by the ?-CD molecules. In (b), dynamic rheology confirms 
this hypothesis as a hm-chitosan (0.25%)/blood (50%) displays gel-like characteristics 
(G? [closed red circles] and G?[closed green triangles]), whereas a hm-chitosan 
(0.25%)/blood (50%)/?-CD (3%) mixture displays the rheology of a freely-flowing liquid 
(G? [open red circles] and G?[open green triangles]).                                                     121 
 
Figure 7.1. Photolysis of MGC, a photobase generator (PBG). Upon UV irradiation, the 
molecule dissociates and releases OH?.                                                                           127                                                                       
 
 
 
 
 
 
 
 
 
 1 
Chapter 1 
Introduction and Overview 
 
 
1.1 Problem Description and Motivation 
 
. 
In the current age of micro- and nanotechnology, there is a great deal of interest in 
making structures, machines, and robots at smaller scales.3 While processing techniques 
for metals or plastics are well-established at the macroscale (e.g., > 10 cm), the same 
techniques can be difficult to implement at smaller scales. For example, the machining or 
molding of a small (~ 5?10 mm) robot made of metal or plastic is not an easy task. This 
problem has an important context particularly within the field of nanomedicine. The 
popularized vision of nanomedicine is to create nano-sized robots, ?nano-bots,? which 
patrol through the body, responding to environmental cues to repair tissue and cure 
disease when and where required.4 Despite the fact that such an intriguing and ambitious 
goal has driven an enormous amount of research over the past 20 years,5 it is important to 
take a step back and carefully consider some of the key points Edward Purcell made in 
his popular 1977 article ?Life at Low Reynold?s Number.?6 Purcell emphasized that 
everything at the nano-scale essential becomes ?sticky,? and, as such, it is very difficult 
to design hard nanoscale mechanical structures which move and function in a manner 
uniquely independent from Brownian motion. In plainer terms, building a robot is hard 
enough?.building a robot to operate within a world of sticky, slimy, and grimy ?muck?  
approaches the impossible.  
 2 
In this context, it is useful to look to nature for inspiration to our nano-machine 
blueprints, as opposed to simply down-sizing successful macro- or micro-mechanical 
designs. Nature?s equivalents to crawling mini-robots are creatures such as geckos, 
earthworms, and ants. It is worth noting that these creatures are generally ?soft?, i.e., they 
are not hard like metal or plastic, but gooey and pliable like jello or silly putty. Rather 
than building hard nano-bots to achieve a rigid endpoint, a more sensible approach would 
be to learn the inner-workings and conditions of these soft biological systems and, 
subsequently, build structures which complement or cleverly manipulate them in 
therapeutically beneficial ways.  
An excellent system to first study as a means to improved nanomaterial design is 
the biological cell. No machinery in the history of mankind has ever come close to 
rivaling its complex set of functionalities and wonderfully robust design. The cell is truly 
a ?soft machine? which is able to perform such tasks as rapidly build AND dissemble rods 
and trusses (actin filaments, intermediate filaments, microtubules), periodically replenish 
its outer shine (synthesis and transport of phospholipids to cell membrane), and reliably 
respond to myriad electrical and molecular signals from the external environment 
(Na+/K+ membrane-associated ion channels). These cellular processes occur via self-
 assembly (i.e. they are thermodynamically driven, ?G < 0 ) with in  a soft environment. 
Note that from a materials characterization standpoint, the cell is not a bag of water, but 
rather a lump of jello. Understanding how the cell?s machination works within this 
?squishy? and ?sticky? environment is key to making progress in almost any given sub-
 discipline of nanomedicine including targeted delivery of drugs or genes, controlled 
release, tissue repair and engineering, and wound healing. 
 3 
1.2 Proposed Approach 
We take inspiration from biology to build self-assembled materials which are able 
function as soft machines for improved therapies in nanomedicine. In this dissertation, we 
explore the integration of polymers with vesicles as a route to creating new and useful 
classes of soft materials. Vesicles, of course, are reminiscent of cells in a few key ways: 
(1) they have a very similar bilayer structure to cell membranes, (2) they have an aqueous 
core which can hold proteins and/or small molecules and (3) they are able to split/fuse 
based on particular environmental cues.7-8 Additionally, the cell uses actual nano-scale 
~100 nm vesicles to transport molecules into and out of the cell. The polymers used in 
our studies are reminiscent of many of the structural polymers which interact with cell 
membranes and assist in the assembly and transport of vesicles within cells. 
Using these 2 key ingredients, (1) vesicles and (2) biopolymers, we blueprint 
self-assembled systems in which the polymer plays an active role in creating a carrier or 
matrix for the vesicles. The resulting hybrid structures impart increased stability, or 
functional restraint of the self-assembled vesicles. Conversely, the vesicles impart 
enhanced functionality to the biopolymer. Such ?easy? approaches to building materials 
involve dialing up conditions and environmental triggers such that vesicles and 
biopolymers build and/or disassemble themselves within aqueous environments. Hence, 
these approaches are not labor intensive. However, we are indeed able to achieve high-
 levels of functionality from these systems simply by understanding and appropriately 
manipulating the factors which drive key self-assembly events. Four specific classes of 
such systems are investigated. These are depicted in Figure 1.1 and briefly described 
below. 
 4 
 
 
Figure 1.1. Overview of the Four Blueprinted Self-Assembled Soft Systems. (a) 
Controlled-Release Jello: pH-Sensitive Vesicles within Biopolymer Gels, (b) 
?Mothership? Drug Carriers: Biopolymer Capsules Containing Vesicles; (c) Band-Aids 
for Chronic Wounds: Biopolymer Films Capturing Vesicles; (d) Self-Assembled 
Hemostats: Associating Biopolymers Gelling Vesicles/Cells 
 
 5 
1.2.1 pH-Sensitive Vesicles within Biopoylmer Gels 
Vesicles which undergo a self-assembled transition into micelles upon increase in pH will 
be embedded within a gelatin gel. Here, we anticipate that a biopolymer gel network will 
form ?around? a solution of vesicles, without disrupting the vesicles (Figure 1.1a). The 
presence of intact vesicles would presumably enhance the functionality of the gel by 
protecting encapsulated molecules and providing a sustained release of model drug. 
Presence of intact vesicles within the gel will be tested by a few different experimental 
methods. Next, a high pH-buffer solution will be placed in contact with the gel. As the 
high pH buffer diffuses into the gel, we will track the movement of a vesicle-micelle 
front through the gel as a function of time, and we will also explore the types of vesicle-
 micelle patterns created by varying the gel geometry and the buffer-gel contact points. 
Lastly, we will attempt to characterize the vesicle-to-micelle transitions within the gel by 
packaging dye molecules within the vesicles and attempting to use pH as a trigger for 
their release.  The experimental design and complementing results for this system are 
explained in detail in Chapter 3. 
 
1.2.2 Vesicles Loaded into Biopolymer Capsules 
Next, we plan to entrap vesicles inside microcapsules made from biopolymers. We call 
these structures ?motherships? as smaller containers, i.e. ?babyships,? are protected 
within a larger container until degradation of the mothership capsule shell. Motherships 
are formed when droplets of a cationic biopolymer solution mixed with vesicles are 
added to a solution of anionic biopolymer or surfactant.5-7 A physical crosslinking occurs 
at the interface of the drop, leading to an interfacial shell (Figure 1.1b) and this shell 
 6 
protects the contents of the drop from external stimuli. Capsule size is determined solely 
by the size of the drop and can be varied from a few millimeters to a few microns. Our 
goal is to trigger the release of vesicles by tailoring the capsule surface to degrade in 
response to specific pH transitions. Also, we hope to develop structures displaying 
features on hierarchy of length scales (from macro to micro to nano) using this single-
 step drop method by placing micro-scale mothership capsules containing nano-scale 
vesicles within macro-scale capsules.  Details on the results of this proposed approach are 
explained in Chapter 5.  
 
1.2.3  Vesicles Captured Onto Biopoylmer Films 
A challenging problem in materials science is the ability to anchor ?soft? biomolecular 
nanostructures such as vesicles to ?hard? surfaces such as gold. For our 3rd blueprint, we 
approach a simple method to accomplish this kind of anchoring using an amphiphilic 
biopolymer as an soft ?interconnect?. The polymer is hydrophobically-modified chitosan 
(hm-chitosan), which is obtained by covalently attaching alkyl tails to the backbone of 
chitosan. We plan to electrodeposit films of hm-chitosan onto microscale gold cathodes 
formed by lithography on a silicon wafer. Such hm-chitosan films could potentially 
capture drug-loaded vesicles from solution (Figure 1.1c). If the vesicles remain strongly 
bound to the films, a method of therapeutically functionalizing the film with multiple 
agents which are protected from degradation would be achieved in a quick one-step 
process. We envision such films being used for protection and controlled release of drug 
to accelerate the healing of chronic wounds of diabetics and burn victims. Additionally, 
 7 
the immobilized vesicles could also be use for biosensing applications. The results of this 
approach in explained in Chapter 5.     
 
1.2.4 Gelling Cells  with Biopolymers 
 
Lastly, we blueprint a system involving biopolymers and cells, instead of vesicles. The 
ability to rapidly self-assemble into a flow-impeding barrier on demand is an ideal feature 
of hemostat, i.e. a material which controls hemorrhage. Indeed, the self-assembly of 
fibrinogen into fibrin, the hemostatic ?plug? which is naturally observed in mammalian 
organisms in response to bleeds, is a rapid process regulated by a highly nuanced cascade 
of previous events. Here, we aim to achieve an equally effective set of coordinated via 
interactions between hm-chitosan and the blood hematocrit. We anticipate based on 
previous work done in our lab with vesicles the biopolymer will form an ?artificial clot,? 
via interaction of hydrophobes and the hydrophobic bilayers of the blood cells (Figure 
1.1d). In this way, the biopolymers would act as bridges between adjacent cells, thus 
forming an elastic 3-dimensional network with the cells themselves acting as physical 
crosslinks. Additionally, we plan to utilize the amphiphilic supramolecule, ?-cyclodextrin 
(CD), which is a barrel-shaped molecule featuring a hydrophobic interior, so as screen 
the hydrophobic interactions and, in this way, reverse the mechanism of gelation. 
Detailed explanations of the designed experimental and results on such ?reversible 
hemostats? are given in Chapter 6. 
 
 
 8 
1.3 Significance of this Work 
  
The studies described in this dissertation are potentially significant from two different 
standpoints: (a) they provide fundamental insight into self-assembly processes, such as in 
biological systems; and (b) the studies provide a foundation for new therapeutic concepts 
in the field of nanomedicine. 
 
Firstly, from a scientific standpoint, the interactions between vesicles and 
biopolymers are crucial to the behavior of cell membranes and membrane-bound 
organelles within a cell. As is well known, every biological membrane consists of a 
combination of lipids as well as biopolymers (proteins or polysaccharides) either 
spanning the membrane or tethered on one side of the membrane. Our studies on cells 
mixed with amphiphilic biopolymers presented in Chapter 6 can certainly provide some 
immediate ?hands-on? insights into these sorts of interactions. Additionally, our 
controlled release studies in Chapters 3 (gels) and 4 (capsules) can potentially give 
indirect information on vesicle-polymer interaction at the nano-scale. Chapter 4 gives 
some important scientific insights into building hierarchical structures via self-assembly 
which have interesting implications for biomimetic organelles, cells or tissues. 
Furthermore, the results shown on the interactions between ?soft? vesicles and ?hard? 
surfaces as discussed in Chapter 5 is of fundamental importance in the physical sciences.  
 
From the standpoint of utility, all four systems display either proof-of-concept 
level or immediate application to an existing biomedical need. The vesicle gels and 
 9 
vesicle-loaded capsules described in this dissertation may be useful for controlled release 
applications. Compared to vesicles alone, the above vesicle-bearing hybrid materials 
present some benefits. For starters, the stability of vesicles is enhanced by embedding 
them in the soft biopolymer matrix. Moreover, the presence of an additional transport 
barrier enables a slower and more extended rate of release for molecules encapsulated in 
the vesicles. As an added benefit, encapsulation within vesicles may also help in 
maintaining the bioactivity of drugs and proteins.9 Vesicles may also be useful as storage 
depots for hydrophobic drugs that cannot be loaded otherwise into gels or capsules. 
Finally, we will show that it is relatively easy to confer targeting capabilities (either by 
magnetic fields or via antibodies) to the capsules, in comparison to the vesicles.   
 
The vesicles on biopolymer films may also be useful in controlled release, but 
more specifically for the application to chronic wound treatment. Biopolymer films 
presented in Chapter 5 can be fabricated in a variety of sizes. As such, we can envision a 
larger macro-sized film functionalized with therapeutically-loaded vesicles that can 
completely cover up a chronic wound or burn injury. The vesicles would provide 
protection and sustained release of the therapeutics, while the anti-microbial chitosan film 
both prevents fluid exudation and bacterial invasion. These films would also allow for 
ample oxygen transfer to the injury during administration, which is very important for the 
outcome of the underlying tissue. 
 
Amphiphilic biopolymers for the gelation of cells have an immediate utility in 
hemorrhage control. Chapter 6 presents some important in vitro and in vivo work on these 
 10 
materials for treating bleeding injuries. The material used in those studies, hm-chitosan, 
is very low cost, and it achieves similar efficacies to hemostats 3 orders of magnitude 
higher in cost. Furthermore, the use of cyclodextrin in that work as an ?anti-clotting? 
complement system offers a functionality which no other hemostatic material has 
achieved. The reversibility of this system via self-assembly is highly clinically significant 
for two key reasons. First, it allows for the surgeon treating the patient to easily clean out 
the hemostatic material and identify the injury under controlled circumstances so as to 
plan the optimal regimen of treatment for the patient for full recovery. Secondly, the anti-
 clotting aspect of the system provides a mitigation against unwanted clotting, a problem 
which is a major concern for any new hemostat undergoing clinical trials.  
 
 
 
 
 
 
 
 
 
 
 
 
 
 11 
Chapter 2 
Background 
 
In this chapter, we describe some of the basic properties of vesicles, associating 
polymers, and biopolymers. We then briefly describe the techniques that we will use to 
study these resulting ?soft? systems, specifically: rheology, neutron scattering, and cryo-
 TEM. The aspects discussed here are of a general nature; literature dealing with more 
specific aspects are discussed in the Introduction sections of succeeding Chapters 3-6. 
 
2.1 Vesicles and Liposomes 
Vesicles are self-assembled capsules formed in water by lipids, surfactants, or 
block copolymers.10-11 The molecules that form vesicles are amphiphilic, with a 
hydrophilic head (depicted as a blue sphere in Figure 2.1) and hydrophobic tail(s) (shown 
in red). The shell of the vesicle is a bilayer (ca. 2-5 nm in thickness) of these amphiphilic 
molecules, with the hydrophilic heads on both sides of the bilayer and thereby exposed to 
water, while the hydrophobic tails inside the bilayer are shielded from water. A vesicle 
can be considered to form by the folding of amphiphilic bilayers, as shown in Figure 2.1. 
Vesicles with only a single bilayer (or lamella) are called unilamellar vesicles (ULVs), 
while vesicles with several concentric bilayers are called multilamellar vesicles (MLVs) 
and these are also referred to as ?onions?. 
 12 
 
The folding of bilayers into vesicles tends to occur only when the bilayers are 
present at low concentration; at high concentrations, bilayers form a lamellar phase.10 The 
tendency for bilayers to fold is driven by a desire to minimize contact of the hydrophobes 
with water at the bilayer ends. Also, the formation of many vesicles from a single 
extended bilayer sheet increases the entropy of the system. Nevertheless, it is useful to 
remember that, at equilibrium, the amphiphiles usually exist as a lamellar phase; so, the 
vesicle state is often of limited stability. In other words, given sufficient time, vesicles 
will get disrupted and form a dilute lamellar phase. An important exception to this rule 
exists in the case of mixed surfactants, where vesicles can exist as equilibrium 
structures.12 
 
Vesicles formed from lipids are referred to as ?liposomes?. The term lipid usually 
refers to amphiphiles that have a biological origin and typically, such molecules have two 
hydrophobic (acyl) tails. Lipid bilayers constitute the membranes found at the boundary 
Figure 2.1.  The structure of vesicles formed by the self-assembly of amphiphiles. 
The vesicle is formed by the folding of an amphiphilic bilayer that is about 2-5 nm in 
thickness. 
 13 
of every living cell as well as many intracellular organelles. The classification of lipids is 
done based on their headgroup type ?  for example, phospholipids have a phosphate 
moiety in their headgroup. Among the phospholipids, the phosphatidylcholines or 
lecithins are a common variety. Lipids tend to have a very low solubility in water because 
they have two hydrophobic tails. As a result, an organic solvent is usually employed in 
preparing lipid vesicles.  
 
The tendency of lipids to form bilayers or vesicles can be rationalized from the 
geometry of these molecules. Generally speaking, the role of molecular geometry in 
dictating the self-assembly of amphiphiles can be understood by a term called the critical 
packing parameter or CPP, which is defined as follows: 13  
 tail
 hg
 CPP
 a
 a
 =    (2.1) 
where ahg is the effective area of the amphiphile headgroup and atail is the average area of 
the amphiphilic tail. Amphiphilic molecules having atail ? ahg, i.e., CPP = 1, tend to 
assemble into bilayers or vesicles (Figure 2.2). Note that the shape of these molecules 
resembles that of a cylinder. In contrast, molecules with a larger headgroup area than tail 
tend to favor curved structures, specifically micelles. A CPP of ? corresponds to 
spherical micelles while a CPP of ? corresponds to cylindrical (rodlike or wormlike) 
micelles. 
 
 In addition to lipids (2-tailed amphiphiles), mixtures of single-tailed amphiphiles, 
one cationic and the other anionic, can also form vesicles.12 The formation of such 
?catanionic? vesicles can also be understood via the CPP concept (Figure 2.2). In this 
 14 
case, each individual surfactant molecule resembles a cone because of the electrostatic 
repulsion from its headgroup. When mixed together, however, the cationic and anionic 
headgroups mutually mitigate their repulsive electrostatic effects, leading to a significant 
reduction in headgroup area. The combination of these molecules thus resembles a 
cylinder, and consequently leads to vesicle structures. Interestingly, these surfactant 
vesicles tend to spontaneously form when the two individual surfactants are mixed. 
Moreover, the vesicles are indefinitely stable, which suggests that they may actually be 
equilibrium structures. 
 
=C
 HO O
 C
 O O
 _ -H
 Fatty Acid Surfactants
 water
 water
 Lipids, 2-tailed amphiphiles
  
 
Figure 2.2. Schematic illustration of the formation of vesicles by self-assembly of 
amphiphilic molecules in water. Due to their cylindrical geometry, lipids, which are two-
 tailed amphiphiles, can form vesicles in aqueous solution upon folding of lipid bilayers 
into spheroid structures represented in the cartoon above. Also, pairs of fatty acid 
surfactants can collectively take on a cylindrical geometries and, thus, form vesicles in 
the same way as lipids. 
 
Vesicles can also be formed from single surfactants in some rare cases.14-15 Single 
ionic surfactants usually form micelles, but in the case of fatty acids and their soaps, 
 15 
vesicles can form over a limited pH range around the pKa of the acid.15 Fatty acids are 
among the simplest types of amphiphiles that exist and there is speculation that their self-
 assembly into vesicles could have occurred in pre-biotic earth, eventually culminating in 
the origin of life.14 The mechanism for vesicle formation from fatty acids is not fully 
understood; however, it is significant that vesicles form only at intermediate pH values at 
which the acid is partially ionized. Thus, fatty acid molecules in vesicles tend to exist as 
pairs of ionized and unionized molecules, and presumably such pairs self-assemble into 
structures exhibiting geometries required to form bilayers (see Figure 2.2).       
       
 
2.1.1 Vesicle Preparation from Lipids 
 As mentioned, lipids are insoluble in water and at equilibrium they tend to form a 
lamellar phase. Therefore, preparation of lipid vesicles calls for the use of an organic 
solvent and some input of energy.1,11 First, the lipid(s) are dissolved in an organic solvent 
such as chloroform. Thereafter, the solvent is removed by evaporation to yield a dry lipid 
film. This film is then hydrated by adding water (or a buffer solution) at a temperature 
above the gel-to-liquid crystal transition of the lipid (Figure 2.3a). The solution is gently 
stirred during this process and the result is the formation of large multi-lamellar vesicles 
(MLVs) in solution. To convert the MLVs to unilamellar vesicles (ULVs), the lipid 
solution is either sonicated or extruded through a polycarbonate filter of given pore size. 
Sonication tends to produce small unilamellar vesicles (15 ? 50 nm in diameter), whereas 
extrusion is typically used to produce unilamellar vesicles with a diameter on the order of 
100 nm (Figure 2.3a). 
  
 16 
While the above methods yield nanometer-sized vesicles, it is also possible to 
obtain giant unilamellar vesicles (GUVs, or giant vesicles in short) that are several 
microns in diameter.2 Giant vesicles can be seen directly by optical microscopy, typically 
in phase contrast mode. A popular method to form giant vesicles in the size range 5 ? 200 
?m is by electroformation (Figure 2.3b). This involves application of an AC voltage 
across lipid-coated electrodes spanning a water-filled chamber. In this process, the giant 
vesicles break off from the lipid film and their size is controlled by the AC voltage and 
frequency.  
. 
 17 
Figure 2.3. Preparation of unilamellar lipid vesicles of various sizes: (a) small or 
large vesicles (20 nm ? 200 nm); 1 (b) giant vesicles by electroformation. 2 
(a) 
Silicone 
spacer
 AC
 ITO 
coated  
glasses
 Dry lipid 
film
 (b) 
 18 
2.2 Associating Polymers 
 The term associating polymer refers to a water-soluble polymer that has an 
amphiphilic character.16 Typically, the polymer backbone is hydrophilic, while 
hydrophobic groups are either present at the ends of the chain (this is called a telechelic 
structure) or the hydrophobes are tethered by chains to the polymer backbone (this is 
referred to as a comb-graft structure). Associating polymers have been synthesized by 
attaching hydrophobes to a range of water-soluble polymers, including polyethylene 
oxide (PEO) and polyacrylamide (PAAm) as well as to biopolymers such as cellulose and 
chitosan.16-17 We will work with chitosan-based associating polymers in this study.  
When added to water, hydrophobes on the polymer associate or self-assemble in 
Figure 2.4. Archiecture of a telechelic associating polymer and the structures formed 
by its self-assembly in aqueous solution 
NETWORK 
 19 
much the same way as surfactant hydrophobes.16,18 In analogy to the micelles formed by 
surfactants, polymer association is believed to result in ?flower micelles? (shown in 
Figure 2.4 for the case of the telechelics), with hydrophobes from many chains present at 
the center of these micelles. Note that at low polymer concentrations, there is significant 
intra-polymer association, while at higher concentrations, there is a shift to inter-polymer 
associations. Thus, at high polymer concentrations, the flower micelles function as 
crosslinks in a transient network, thereby enhancing the solution viscosity. For high 
molecular weights, the polymer will both associate as well as entangle with other chains. 
 
2.3 Biopolymers 
Macromolecules of biological origin fall broadly under three classes: polypeptides 
or proteins; polynucleotides; and polysaccharides.19 For the purposes of this study, we 
will focus on polysaccharides. The polysaccharide that is of especial interest to us is 
chitosan and we describe its properties below in more detail. The common theme with the 
chosen biopolymers is their ability to render viscosity to water by forming entangled 
networks or gels.       
 
2.3.1 Gelatin 
Gelatin is protein produced by the partial hydrolysis of collagen extracted from the bones 
and connective tissues of animals such as cattle, pigs and horses. The natural molecular 
bonds between individual collagen strands are broken down into a form that rearranges 
more easily. Gelatin forms a solution of high viscosity in water, which sets to a gel upon 
cooling, and its chemical composition is similar to that of its parent molecule collagen.19 
 20 
The same triple helical Gly-X-Y domains abundant in collagen fibers, where X is 
typically proline and Y is typically hydroxyproline, form the physical crosslinks below 
the phase transition temperature of 32?C which cause gelatin to form an elastic gel in 
water.  
 
2.3.2 Chitosan 
Chitosan is a linear polysaccharide obtained by the deacetylation of chitin.20 
Chitin, in turn, is a natural polysaccharide that constitutes the hard exterior shell of 
insects and crustaceans. Among biological polymers, chitin is next only to cellulose in 
abundance. However, while chitin is insoluble in water, its deacetylated derivative, 
chitosan, is water-soluble under acidic conditions (pH < 6.5). Under these conditions, the 
amine groups along the chitosan backbone are ionized and chitosan acts as a cationic 
polyelectrolyte. Note that chitosan is strictly a copolymer of mostly D-glucosamine (?-
 (1,4)-2-deoxy-2-amino-D-glucopyranose) sugars and a few of the N-acetyl-D-
 glucosamine (?-(1,4)-2-deoxy-2-acetamido-D-glucopyranose) sugars from the parent 
chitin. The structures of these sugars are shown in Figure 2.5. 
 21 
 
Current scientific and technological interest in chitosan is motivated by a number 
of factors. 20 First, chitosan is a biocompatible and biodegradable polymer. It is one of the 
few cationic biopolymers, and as a result, it can interact with anionic cell membranes. 
Second, chitosan confers anti-bacterial properties to substrates. As a result, chitosan finds 
application in tissue regeneration, artificial skin constructs, wound dressings and sutures, 
drug delivery, antibacterial coatings, and bioseparation membranes.20-24 Third, there is an  
 
Figure 2.5. Structures of the parent sugars in (a) chitin and (b) chitosan. The N-
 acetyl-D-glucosamine sugar in chitin is deacetylated to give the D-glucosamine sugar 
in chitosan. 
H
 H
 H
 OH
 O
 OH
 NH2
 CH2OH
 n
  
H
 H
 H
 OH
 O
 OH
 NH
 CH2OH
 O
 n
  
(a) (b) 
 22 
environmental benefit to using chitosan since the parent chitin is usually obtained 
from food-processing wastes (e.g., crab, shrimp or lobster shells). For these reasons, there 
is ample interest in chitosan and many researchers have also attempted to modify the 
parent polymer to confer it unique properties. 22,25-27 One such modification is to attach 
hydrophobic groups to chitosan, and this is discussed below.  
 
 
2.3.3 Chitosan Modification 
Chitosan can be modified easily due to its amine groups as well as its primary and 
secondary hydroxyls. Modifications have been done to improve chitosan solubility in 
water, to increase its chelating ability, and to modify its antibacterial effect.17,22,28 Our 
particular interest is in attaching hydrophobic groups to the chitosan backbone to alter its 
association behavior in water. The synthesis of hydrophobically-modified chitosan 
(hm-chitosan) is rather straightforward and can be performed under mild conditions.17 
The procedure involves reacting the chitosan with an n-alkyl aldehyde and a typical 
recipe is described below. In the process, the amine (NH2) groups are converted into 
O
 NH2H H
 OH
 H OH
 O
 NH2
 H
 H H
 OH
 OH
 OO
 O
 H
 HNH
 OH
 H
 OH
 C
 12
 H
 25
 Figure 2.6.  Structure of hydrophobically-modified chitosan (hm-chitosan) with C12 
hydrophobic tails. 
 23 
NH-R groups, where R is the n-alkyl moiety. The structure of hm-chitosan containing C12 
hydrophobic tails is shown in Figure 2.6. Note that this is a comb-graft type of 
associating polymer. 
 
A typical procedure for synthesizing hm-chitosan with C12 tails involves the 
following steps.17-18 First, n-dodecyl aldehyde is added to an acidic chitosan solution in a 
water-ethanol mixture, followed by addition of sodium cyanoborohydride. The molar 
ratio of aldehyde to chitosan monomer(s) is fixed at a certain value (e.g. 2.5%). The 
reaction yields the hm-chitosan, which is then precipitated by raising the pH and adding 
ethanol. Next, the precipitate is purified by washing with ethanol followed by deionized 
water. The final hm-chitosan precipitate is re-dissolved in acetic acid solution and the 
concentration is recalibrated. This solution tends to be viscous due to associations 
between the hydrophobes (this is a qualitative indication that the synthesis has been 
successful). The degree of hydrophobic substitution in the final product can be compared 
to the value expected from stoichiometry using 1H NMR.  
 
Hydrophobically modified chitosans with n-alkyl pendant chains can also be 
synthesized by alternate routes, e.g., by reacting with alkyl carboxylic acids,29 or alkyl 
acid anhydrides,30 or alkyl acid chlorides.31 High substitution levels (> 10%) have been 
reported via the acid chloride method. These alternate routes are not attempted in the 
present study. 
 
 
 24 
2.3.4 Gellan Gum 
Gellan Gum is a polysaccharide produced extracellularly by a microorganism 
(Pseudemonas elodea); it is a straight-chain heteropolysaccharide and it is constituted of 
a repeating unit of four monosaccharide molecules, i.e. glucose, glucuronic acid, glucose, 
and rhamnose, thus have one carboxyl group in the repeating unit. This carboxyl group 
allows the biopolymer to carry a negative charge in aqueous media. A schematic of the 
biopolymer is shown below: 
 
 
Figure 2.7. Structure of Gellan Gum 
 
 
 
 
 
 
 
 
 
 
 25 
2.4 Cyclodextrins 
 
 
 
 
 
 
 
Figure 2.8. Truncated cone-shaped conformation of ?-CD.32 
 Cyclodextrins (CDs) are cyclic oligosaccharides containing D-(+) glucopyranose 
units attached by ?-(1,4) glucosidic bonds, as shown in Figure 2.7.33They are rigid, 
truncated cone-shaped structures, with an internal cavity of size 5 to 8 ? depending upon 
the number of glucopyranose units. The wide side of the truncated cone is bordered by 
the secondary hydroxyl groups (2-OH and 3-OH), while the primary hydroxyl groups 
(6-OH) are on the narrow side.  The molecule is stiffened by hydrogen bonding between 
the 2-OH and 3-OH groups around its outer rim. Note that all hydroxyl groups are located 
on the outside of the molecular cavity, thereby making the outer surface hydrophilic. On 
the other hand, no hydroxyl groups are located in the inner cavity, which is thus 
hydrophobic. CDs thus have hydrophilic outer surfaces and hydrophobic inner cavities. 
Because of their unique structure, CDs can form host-guest inclusion complexes with 
various hydrophobic guest molecules or hydrophobic parts of these molecules.32-38 Note 
 26 
that the bonding between the CD and the guest is through non-covalent interactions. 
Table 2.1 lists the properties of the three naturally occurring CDs, which are labeled ?-, 
?-, and ?-CDs.39 These natural CDs are produced from starch by enzymatic degradation. 
In our research, we will use CDs to create new TR fluids, as described in Chapter 5. 
 
Type of CD Number of 
glucose units 
Cavity diameter 
? 
Molecular 
Weight 
Solubility in 
water (g/L) 
? 6 4.7-5.3 972 145 
? 7 6-6.5 1135 18.5 
? 8 7.5-8.3 1297 232 
 
Table 2.1. Properties of the three naturally occurring cyclodextrins. 
 
 
 
 
 
 
 
 
 
 
 
 
 27 
2.5 Characterization Technique ? I: Rheology 
 
Rheology is formally defined as the study of flow and deformation in materials.40 
Rheological measurements provide important information on soft materials, specifically 
on the relation between microstructure and macroscopic properties. These measurements 
are typically performed on a rheometer (Figure 2.9) under steady or dynamic oscillatory 
shear. Typical geometries used in rheometers are the cone-and-plate, the parallel plate, 
and the concentric cylinder or Couette. 
 
Figure 2.9.  Photograph of the Rheometer RDA-III strain-controlled rheometer being 
operated in a cone-and-plate geometry. 
 28 
In steady shear rheology, the sample is subjected to a constant shear-rate ??  (e.g. 
by applying a continuous rotation at a fixed rate on a rotational instrument), and the 
response is measured as a shear-stress ?. The ratio of shear-stress ? to shear-rate ??  is the 
(apparent) viscosity ?. A plot of the viscosity vs. shear-rate ??  is called the flow curve of 
the material.  
 
In dynamic or oscillatory rheology, a sinusoidal strain 0 sin( )t? ? ?= is imposed 
on the sample. Here, ?0 is the strain-amplitude (i.e. the maximum applied deformation) 
and ? is the frequency of the oscillations. The sample response will be in the form of a 
sinusoidal stress 0 sin( )t? ? ? ?= +  which will be shifted by a phase angle ? with respect 
to the strain waveform. Using trigonometric identities, this stress waveform can be 
decomposed into two components, one in-phase with the strain and the other out-of-phase 
by 90?: 
 0 0sin( ) cos( )G t G t? ? ? ? ?? ??= +   (2.2) 
  where G?  = Elastic or Storage Modulus   
 and G? = Viscous or Loss Modulus                 
 
 The elastic modulus G? is the in-phase component and provides information about 
the elastic nature of the material. Since elastic behavior implies the storage of 
deformational energy, this parameter is also called the storage modulus. The viscous 
modulus G?, on the other hand, is the out-of-phase component and characterizes the 
viscous nature of the material. Since viscous deformation results in the dissipation of 
 29 
energy, G? is also called the loss modulus. For these properties to be meaningful, the 
dynamic rheological measurements must be made in the ?linear viscoelastic? (LVE) 
regime of the sample. This means that the stress must be linearly proportional to the 
imposed strain (i.e., moduli independent of strain amplitude). In that case, the elastic and 
viscous moduli are only functions of the frequency of oscillations ?, and are true material 
functions. A log-log plot of the moduli vs. frequency, i.e. G?(?) and G?(?), is called the 
frequency spectrum or dynamic mechanical spectrum of the material. Such a plot 
represents a signature of the material microstructure.  
 
 The important advantage of dynamic shear is that it allows us to characterize 
microstructures without disrupting them in the process. The net deformation imposed on 
the sample is minimal because the experiments are restricted to small strain amplitudes 
within the LVE regime of the sample. As a result, the linear viscoelastic moduli reflect 
the microstructures present in the sample at rest. This is to be contrasted with steady 
shear, where the material functions are always obtained under flow conditions 
corresponding to relatively drastic deformations. We can therefore correlate dynamic 
rheological parameters to static microstructures, and parameters under steady shear to 
flow-induced changes in microstructure. 
 
 
 
 
 30 
2.6 Characterization Technique ? II:  
      Small Angle Neutron Scattering (SANS) 
 
Scattering techniques are invaluable probes of the micro- and nanostructure in 
soft materials.41 The basic principle underlying all scattering techniques is that the 
intensity of scattered radiation is a function of the size, shape, and interactions of the 
?particles? present. For aqueous systems, small-angle neutron scattering (SANS) is the 
technique of choice because contrast between the ?particles? and the solvent can be easily 
achieved by switching H2O with D2O. Also, the incident radiation in SANS is composed 
of neutrons having a wavelength ~ 7 ?, and as a result, SANS is useful in probing size 
scales on the order of a few nm. SANS experiments require a nuclear reactor to generate 
neutrons and we are fortunate to have one of the premier SANS facilities in the world 
close to UMD at NIST in Gaithersburg, MD.  
 
The basic geometry of a SANS experiment is illustrated in Figure 2.10. Neutrons 
emitted from a nuclear reactor are selected at a particular wavelength and wavelength 
spread using a velocity selector, collimated by several lenses, and passed through a 
sample chamber. The neutrons scattered by the sample are collected on a 2-D detector. 
This 2-D data is corrected and placed on an absolute scale using calibration standards. It 
Figure 2.10.  Schematic of a SANS experiment. 
? 
 31 
is then converted into a plot of scattered intensity I vs. scattering or wave vector q by 
spherical averaging. The wave vector q is related to the scattering angle and wavelength 
by: 41  
 
4
 sin
 2
 q
 ? ?
 ?
 ? ?= ? ?
 ? ?
  (2.3) 
Here, ?  is the wavelength of the incident radiation and ?  is the scattering angle. Thus, q 
can be considered an inverse length scale, with high q corresponding to small structures, 
and vice versa.  
 
 The SANS intensity I(q) from a structured fluid containing np particles per unit 
volume can be expressed in the following manner:41 
 p( ) ( ) ( )I q n P q S q= ? ?  (2.4) 
where P(q) is called the form factor and S(q) the structure factor. P(q) is the scattering 
that arises from intraparticle interference, which is a function of the particle size and 
shape. S(q) arises from interparticle interactions and thereby reflects the spatial 
arrangement of particles in the sample. When the particles are in dilute solution or are 
non-interacting, the structure factor S(q) ? 1 and the SANS intensity I(q) can then be 
modeled purely in terms of the form factor P(q). Different expressions exist for the form 
factor P(q) for various particle geometries. By fitting the appropriate P(q) to the SANS 
data, one can obtain the characteristic sizes of the particles present. 
 
 
 
 
 32 
2.7 Characterization Technique ? III: Dynamic Light Scattering (DLS) 
 
 Static scattering techniques such as SANS provide important information about 
the quiescent structure in complex fluids. Dynamic scattering techniques have a  
complementary role in that they probe structural relaxations and dynamics. In particular, 
dynamic light scattering (DLS) probes the Brownian motion of particles in the fluid. This 
method can give a reliable estimate of particle size under certain limiting conditions. In a 
DLS experiment, the fluctuating intensity of light scattered from the sample is recorded at 
a certain angle ?. The fluctuations are then correlated to yield the intensity 
autocorrelation function g(2)(q,?) vs. the correlation time ?:42 
  (2)
 2
 ( , ) ( , )
 ( , )
 ( , )
 I q t I q t
 g q
 I q t
 ?
 ?
 +
 =  (2.5) 
Note that in light scattering, the definition of the wave vector is slightly modified as: 
 
4
 sin
 2
 n
 q
 ? ?
 ?
 ? ?= ? ?
 ? ?
  (2.6) 
where n is the refractive index of the medium. The relevance of q in DLS is that 
structural relaxations are probed over length scales on the order of q?1.  
 
The measured intensity autocorrelation function g(2)(q, ?) can be converted into an 
electric field autocorrelation function g(1)(q, ?) through the Siegert relation: 
  
2(2) (1)( , ) 1 ( , )g q f g q? ?= +  (2.7) 
Here, f is an adjustable parameter called the coherence factor that depends on the 
instrument geometry. For a dilute solution of monodisperse spherical particles, the 
 33 
electric-field autocorrelation function is a single exponential whose time decay is 
determined by the translational diffusion coefficient of the particle D: 
  ( )(1) 2( , ) expg q Dq? ?= ?  (2.8) 
From the measured diffusion coefficient, the particle size can be obtained by the Stokes-
 Einstein equation: 
  B
 h6
 k T
 D
 R??
 =  (2.9) 
where kB is the Boltzmann constant, T the absolute temperature and ? the viscosity of the 
solvent (assumed to be a Newtonian liquid). The size obtained from DLS is the 
hydrodynamic radius Rh. The hydrodynamic size is the bare particle size along with any 
solvation layer.    
 
2.8 Characterization Technique ? IV:  
      Cryo-Transmission Electron Microscopy (Cryo-TEM) 
 
 While scattering techniques provide indirect information about the nanostructure 
in a sample, an alternate technique that would allow direct visualization of the structure 
in real space would be extremely useful. Transmission electron microscopy (TEM) can 
potentially reveal structural detail with sub-nanometer resolution. However, the use of 
TEM for self-assembled fluids and soft materials is problematic. For imaging under TEM, 
the solvent must be completely removed, and the process of solvent removal can alter or 
destroy fragile structures such as micelles and vesicles. Also, to achieve contrast, 
structures usually have to be stained with heavy metal salts ? again, the staining process 
might degrade the structures present.  
 34 
 
 The above deficiencies of conventional TEM techniques have led researchers to 
develop an alternative that is particularly suited for aqueous nanostructured fluids. This 
technique is called cryo-TEM, and it involves the rapid freezing of the aqueous sample 
such that the water is vitrified instead of forming ice crystals. In the process, the 
structural details are preserved, and the sample can be imaged under conventional TEM 
at low electron doses. Sample preparation for cryo-TEM is conducted using a controlled  
environment vitrification system (CEVS), illustrated in Figure 2.11. In the CEVS, the 
sample is equilibrated at conditions of controlled temperature and humidity prior to 
plunge vitrification. First, a drop of the sample is placed on a holey carbon film supported 
on a TEM grid. A filter paper is then used to blot the drop, so as to create a thin film of 
the sample spanning the grid holes. The grid is then plunged into the cryogen, liquid 
ethane, thereby rapidly vitrifying the sample. Subsequently, the grid has to be transferred 
to the electron microscope in a dedicated cold stage. Typically, the grid is maintained at  
?170?C at all times, to ensure that there is no formation of ice crystals nor condensation 
of atmospheric water.  
As with any other technique, care has to be taken in interpreting cryo-TEM data. 
One frequent source of artifacts is that the sample blotting process (used to spread a thin 
film) involves a substantial amount of shear. This shear may distort the structures present, 
and one has to take this into account while interpreting images. Despite these artifactual 
concerns, cryo-TEM has now developed into a powerful tool for directly probing the 
structures of various complex fluids. A variety of self-assembled structures including 
micelles, vesicles and liquid crystalline phases have been successfully imaged by cryo-
 35 
TEM. Also, the use of cryo-TEM in biology has provided unique insights into 
phenomena such as endocytosis and vesicle fusion. 
 
 
 
 
 
Figure 2.11.  Photograph of the controlled environment vitrification system (CEVS) 
used for sample preparation in cryo-TEM. The schematics on the left show the 
various steps in the process. 
 36 
Chapter 3 
Biopolymer Gels Containing pH-Sensitive Vesicles  
 
The results presented in this chapter have been published in the following journal article: 
Matthew Dowling, Jae-Ho Lee, and Srinivasa R. Raghavan, ?pH-Responsive Jello: 
Gelatin Gels Containing Fatty Acid Vesicles?Langmuir, 25 (15)
  
, 8519-8525 (2009). 
3.1 Introduction 
 
In recent years, there has been immense interest in creating smart materials, i.e., 
materials that can change their properties in response to external stimuli such as light, 
temperature, pH, or biological targets.43-46 In particular, stimuli-responsive polymer 
hydrogels have attracted much attention, and a stimulus of interest in this context has 
been pH.43-45 Polymers having ionizable groups (acid or base) tend to form gels with 
pH-responsive swelling properties. Such gels generally tend to be swollen when the 
groups are ionized, while they revert to a collapsed or shrunken state when the same 
groups lose their charge. Examples include gels of poly(acrylic acid), poly(methacrylic 
acid) etc. and their copolymers, and these have proven attractive for controlled release 
and drug delivery applications.43-45   
 
An alternate way to impart pH-dependent properties to polymer gels is by 
embedding pH-responsive nanoparticles or nanostructures in their interior.46 In this 
scenario, the polymeric framework itself is unaffected by pH (e.g., there is no change in 
the degree of gel swelling with pH). This approach may be attractive for a variety of 
 37 
reasons. For instance, one may wish to modulate the release of a drug from a gel while 
maintaining a constant gel volume. Secondly, by embedding nanostructures, 
pH-responsiveness can be conferred to materials that by themselves do not have such 
properties. For example, consider hydrogels of gelatin and poly(N-isopropyl acrylamide) 
(NIPAAm).44,47 Both gels have thermoresponsive properties: gelatin gels melt upon 
heating while NIPAAm gels shrink when heated beyond a critical temperature. While 
these polymers are workhorses in many applications, they are not pH-responsive (at least 
not at moderate pH values). One could modify the chemistry of these polymers to make 
the gels pH-sensitive, but that is generally a tedious process. The addition of 
nanostructures to impart pH-responsiveness is generally a much simpler alternative.   
 
In this study, we explore the addition of pH-responsive vesicles into gelatin gels 
and investigate the resulting vesicle-gel hybrids. Unlike ?hard? nanoparticles, vesicles (or 
liposomes) are ?soft? self-assembled structures formed from lipids or surfactants.48 
Unilamellar vesicles of ca. 100 nm diameter can be essentially considered as 
nanocontainers, with a bilayer membrane enclosing an aqueous core.48-50 Vesicles thus 
have the ability to encapsulate hydrophilic solutes in their core, and for this reason they 
are of great interest for drug delivery applications.48-50 In the present context, we are 
interested in embedding vesicles within a polymer hydrogel network. Vesicle-gel hybrids 
have been investigated in the past by several researchers.51-56 The motivation for past 
studies has been the use of these hybrid materials in tissue engineering or drug delivery. 
For example, drug delivery from a vesicle-loaded gel has been shown to occur in a 
prolonged manner compared to that from either the vesicles themselves or the bare gel 
 38 
alone.51-56 To our knowledge though, pH-responsive vesicles have not been combined 
with hydrogels previously ? this is the first study to explore such a combination.            
 
The pH-responsive vesicles we have chosen to examine are those made from 
sodium oleate (NaOA), the sodium salt of a single-tailed fatty acid, oleic acid. It has been 
known for many years that fatty acids can self-assemble into unilamellar vesicles at pH 
values close to their pKa.14-15,57-59 Fatty acid vesicles have been investigated particularly 
by researchers in the field of prebiotic chemistry ? these researchers believe that fatty 
acids and by extension their vesicles may have been components in the prebiotic soup of 
ancient earth (indeed, fatty acids have even been isolated from meteorites).14,58-59 In the 
present study, we exploit the well-known pH-dependence of fatty acid self-assembly.15,57  
For example, NaOA forms vesicles at a pH around its pKa of 8.3, but these vesicles 
transform into spherical micelles at pH values much higher than the pKa, e.g., at pH > 10 
(see the phase diagram in Figure 1).57 We were interested to see whether such a pH-
 induced vesicle-to-micelle transition would occur even if the NaOA vesicles were 
entrapped within a gel. As the results in this paper show, this is indeed the case. The 
above concept can be extended to other pH-dependent vesicles (e.g., those formed from 
certain lipids60). Also, instead of gelatin, a variety of polymer and biopolymer-based 
hydrogels can also be used as the matrix for the vesicles.45,61 
 
Finally, it is worthwhile to discuss some applications for gels loaded with pH-
 responsive vesicles. An obvious one would be in pH-controlled release of hydrophilic 
solutes.43-45 Consider solutes encapsulated within the vesicles, which in turn are loaded 
 39 
into a gel. If such a gel is immersed in an aqueous bath, the solute will slowly diffuse out 
of the gel. We can then use the pH in the bath to modulate the solute release rate. For 
example, at a pH corresponding to intact vesicles, the solute will face two barriers to 
transport, one from the vesicle bilayer and the other from the gel matrix.51-56 On the other 
hand, consider a pH at which the vesicles are disrupted into micelles: in this case, the 
solute will only face resistance from the gel matrix and should therefore be able to diffuse 
out faster. In our studies, we have tested the above concept using a dye, calcein, as a 
model solute, and our findings confirm a pH-tunable release rate.         
 
A vesicle-loaded gel could also serve as a simplistic model for a functional organ, 
with the vesicles as the cellular component and the gel as the extracellular matrix.45,62 
The overall functionality of an organ depends on the ability of cells to communicate with 
neighboring cells.62 Cells of different types may be localized at different parts of the 
organ,  e.g. they may be organized into distinct layers, with their location being strongly 
correlated with cellular function. In this context, it is useful to be able to dictate the 
spatial organization of vesicles within a gel, i.e., to have only certain regions filled with 
vesicles while adjoining regions would not contain any. We show that pH-responsive 
vesicles offer one way to accomplish such a design: specifically, we demonstrate how 
vesicle-to-micelle transitions can be achieved in localized regions within a gel. This 
opens up the prospect of creating a ?biomimetic organ? by placing pockets of vesicles 
loaded with specific molecules at prescribed locations within a gel matrix.   
 
 
 40 
3.2 Experimental Section 
Materials. Sodium oleate (NaOA) (note: oleate = C18 chain with a 9-cis unsaturation) 
was purchased from TCI. Gelatin (Type A, from porcine skin, ~ 300 Bloom), calcein, 
Triton-X100 and Tris-base were purchased from Sigma. Carbonate buffer solution 
(potassium carbonate-potassium borate-potassium hydroxide) corresponding to a pH of 
10 was purchased from Fisher Scientific. All experiments were performed using 
deionized (DI) water. 
 
Preparation of Vesicles and Vesicle-Loaded Gels. 1 wt% (32.8 mM) NaOA was 
dissolved in DI water by heating for 2-3 h at 65?C. The pH of the solution was then 
adjusted to ~ 8.5 by dropwise addition of 1 M HCl. In order to generate unilamellar 
vesicles of consistent size, the vesicle solution was subjected to 5 freeze-thaw cycles, 
followed by passing the sample 10 times through a 100 nm polycarbonate membrane 
using the Mini-extruder? (Avanti Lipids). To prepare vesicle-loaded gels, an NaOA 
vesicle solution of given concentration was combined with a warm (50?C) solution of 
gelatin, with the overall pH adjusted to 8.5 using 1 M NaOH. The resulting mixtures were 
cooled to room temperature to form the vesicle-loaded gels. Further details are discussed 
in the Results section.  
 
Dynamic Light Scattering (DLS). DLS was used to characterize the sizes of vesicles in 
solution. A Photocor-FC light scattering instrument with a 5 mW laser source at 633 nm 
was used at a scattering angle of 90?. A logarithmic correlator was used to measure the 
 41 
intensity autocorrelation function. The hydrodynamic size of the vesicles was extracted 
from the data using the Stokes-Einstein equation.   
 
SANS. SANS measurements were made on the NG-3 (30 m) beamline at the National 
Institutes of Standards and Technology (NIST) in Gaithersburg, MD. Samples were 
prepared in D2O and studied at 25?C in 2 mm quartz cells. The scattering spectra were 
corrected and placed on an absolute scale using calibration standards provided by NIST. 
Data are shown for the absolute intensity I versus the scattering vector q = (4?/?)sin(?/2), 
where ? is the wavelength of incident neutrons and ? is the scattering angle.  
 
Controlled Release Experiments. NaOA vesicles were combined with 15 mM calcein 
dye. To separate unencapsulated dye, the vesicle-dye mixture was passed through a 
Sephadex G-50 size-exclusion chromatography (SEC) column. The vesicle fraction (with 
encapsulated dye) was collected and used in preparing the vesicle-loaded gels. To 
examine the release of dye, 4 g of aqueous buffer solution was added above 6 g of gel in 
the headspace of the containing vials. The concentration of calcein in the buffer was 
measured by UV-Vis spectrometry (Cary Bio 50) as a function of time. Release 
experiments were also done with a control gel (no vesicles). The dye concentration in the 
control gel was kept identical to that in the vesicle-loaded gel.  To determine the latter, 
2 mL of the vesicle fraction from SEC was combined with 50 ?L of 10% Triton-X100 
detergent, thereby disrupting the dye-loaded vesicles and releasing the dye into the bulk 
solution. The dye absorbance (and thereby concentration) could then be measured 
accurately by UV-Vis, and this concentration was used in preparing the control gel.     
 42 
3.3 Results and Discussion 
Creation of Vesicle-Loaded Gelatin Gels. We first describe the creation of vesicle-
 loaded gelatin gels by blending NaOA vesicles and gelatin. We will also address the 
stability of the vesicles within the gels. As mentioned in the Introduction, NaOA vesicles 
are made by adjusting the pH of an NaOA solution. The variation in NaOA solution 
structure as a function of pH is depicted in Figure 3.1, which is an equilibrium titration 
curve. To obtain this data, increasing amounts of 1 M HCl were added to a micellar 
solution of NaOA at pH 10 and the samples were studied by visual observations and light 
scattering. Phase assignments were done in accordance with our observations and these 
are fully in accord with the phase diagram reported previously for oleate solutions by 
Cistola et. al.57 Photographs of selected samples corresponding to different regions in the 
phase diagram are shown in Figure 3.1.  
 
 43 
 
Figure 3.1. Photographs and titration curve for 1% NaOA at 25?C. Increasing amounts of 
1 M HCl are added to a micellar solution of NaOA (pH 10) and the corresponding 
solution pH at equilibrium is shown in the plot. Structural assignments are done in 
accordance with Ref. 10. Photographs of samples corresponding to different regions of 
the plot are shown above. 
 
 
Let us consider the data starting from the origin. At high pH (9.5 or above), 
NaOA forms solutions of spherical micelles and these samples are colorless and 
transparent, as shown by the photograph. As concentrated HCl is added, the solution pH 
drops from about 9.5 to 7.0, a range that spans the pKa of NaOA (which is 8.3). Samples 
in this pH range (interval BC on the plot) are homogeneous and have a strong bluish 
color, as shown by the photograph. These samples are vesicle solutions and the bluish 
 44 
hue is a manifestation of light scattering from the vesicles (Tyndall effect). With further 
decrease in pH, the NaOA samples become unstable and inhomogeneous due to the 
formation of an oil phase (undissociated NaOA). The low pH samples are turbid, milky 
emulsions of the oil and water phases, as seen from the photograph.  
 
The aspect to note from Figure 3.1 is the continuous and reversible transition from 
NaOA micelles at high pH (> 9.5) to NaOA vesicles at moderate pH (~ 8.3). The 
mechanism for this transition (which occurs for all long-chain fatty acids) has been 
discussed in a number of studies and the central factor is believed to be the change in the 
degree of ionization of the fatty acid.15,57 At high pH, the carboxylate groups on NaOA 
are fully ionized and the headgroups of the amphiphile thus bear a strong negative 
charge. The repulsions of these headgroups lead to the formation of spherical micelles. 
Because the micelle size is only around 5 nm, the solutions scatter light weakly and 
therefore appear transparent. When the pH is lowered to a value close to the pKa, 
approximately half the NaOA groups are no longer ionized. The charge may then be 
considered to be shared by two adjacent fatty acid molecules, one ionized and the other 
unionized: i.e., the molecules are effectively paired into dimers.15,57 The net amphiphile 
geometry then becomes conducive to formation of vesicles rather than micelles. The 
vesicles typically range from 50 to 150 nm in diameter (much larger than the micelles), 
and therefore the solutions scatter light strongly, leading to the bluish color.  
 
Having ascertained the pH-sensitive nature of NaOA vesicles, we now proceed to 
discuss their encapsulation within gelatin gels (Figure 3.2). For this purpose, we first 
 45 
made a 1 wt% NaOA solution at a pH of 8.3 and extruded the vesicles through 100 nm 
pores to finally obtain vesicles of an average diameter around 100 nm. We then made a 
solution of 10 wt% gelatin in DI water and adjusted its pH to 8.3. Both the gelatin and 
NaOA solutions were warmed to 50?C and then mixed in a 1:1 ratio by weight. As is 
well-known, gelatin undergoes a sol to gel transition when cooled below 35?C.47 Upon 
cooling to room temperature, we obtained a bluish gelatin gel, as shown by the 
photograph in Figure 3.2c, with an overall concentration of 5 wt% gelatin and 0.5 wt% 
NaOA vesicles. Note that gelatin gels not containing vesicles are colorless (Figure 2a), so 
the bluish color of the gel in Figure 3.2c is a visual indication that vesicles are indeed 
intact in the gel. This is further discussed below. Figure 3.2 depicts the microstructure of 
the vesicle-gel: it consists of vesicles entrapped in a 3-dimensional network of gelatin 
chains. The crosslinks in the gelatin gel are known to be triple helical domains where 
three adjacent chains are linked together.47  
 
Figure 3.2. Photographs and schematics of (a) gelatin gel; (b) NaOA vesicles; (c) gelatin 
gel loaded with NaOA vesicles. The gelatin gel is a 3-D network of gelatin chains, with 
chain segments connected into triple helices at the crosslink points. When vesicles of 
diameter ~ 100 nm are entrapped in the gelatin gel, the initially colorless gel assumes a 
bluish hue due to light scattering from the vesicles. 
 46 
How can we be sure that the vesicles are intact in the gel? One should note that  
stable encapsulation of vesicles in gels has already been demonstrated in a number of 
earlier studies.51-56 In the present case, we cannot do a DLS experiment with the vesicle-
 loaded gel itself because the Stokes-Einstein formulation applies only for vesicles in a 
liquid medium. However, we can put the thermoreversibility of gelatin gels to use here. 
Specifically, we can heat the gelatin gel till it melts (~ to ca. 45?C) and becomes a sol and 
then check for the vesicle size in the sol by DLS. This can be compared with the DLS 
results on an NaOA vesicle solution at 45?C. For 0.5% NaOA vesicles in a 5% gelatin sol 
at 45?C, we measured a mean radius of 46 ? 3 nm from DLS. In comparison, for 0.5% 
NaOA vesicles at the same temperature with no gelatin added, we obtained a similar 
mean radius of 44 ? 1 nm. These results point to the presence of intact NaOA vesicles in 
gelatin gels. Incidentally, both the vesicle-gel and its corresponding sol retain the bluish 
hue of the original vesicle solution.  
 
In addition to DLS, we have also conducted small-angle neutron scattering 
(SANS) experiments at room temperature on both NaOA vesicle solutions and gelatin 
gels containing NaOA vesicles. Figure 3.3 shows a plot of the scattered intensity I vs. 
wave-vector q for two samples: 0.5% NaOA vesicles and the corresponding vesicle gel 
made with 5% gelatin. In both cases, the data follow a slope of ?2 at low to moderate q, 
which is a qualitative indication for the presence of bilayered structures (vesicles).63 By 
plotting the above data on cross-sectional Guinier plots (not shown),64 we can extract the 
thickness of the vesicle bilayer, which is found to be 3.2 ? 0.2 nm in both cases. Further 
modeling of the SANS data to compare vesicle sizes is beyond the scope of this paper 
 47 
because, for the vesicle-gel, one would need to account for the contribution from the 
gelatin network to the scattering intensity. In any case, the SANS data are clearly 
consistent with the presence of vesicles inside the gel. It is also worth mentioning that in 
addition to NaOA vesicles, we have also been able to encapsulate phospholipid-based 
giant unilamellar vesicles (GUVs) in gelatin gels. These vesicles have diameters 
exceeding 10 ?m and so their presence in the gel is easily confirmed using phase-contrast 
and fluorescence microscopy (data not shown).  
 
Figure 3.3.  SANS data at 25?C for 0.5% NaOA vesicles at pH 8.3 (circles) and a 5% 
gelatin gel loaded with 0.5% NaOA vesicles (squares). In both cases, the intensity I 
follows a slope of ?2 at moderate q, which is characteristic of scattering from vesicles.  
 
   
 48 
Inducing Vesicle-Micelle Transitions Within Gels. The above data confirm that NaOA 
vesicles can be entrapped within gels. The next question is whether these vesicles still 
retain their pH-responsive properties. To test the pH response of the vesicle-gels, we 
initially did the following experiment. We placed a 5% gelatin gel loaded with 0.5% 
NaOA vesicles in a vial and filled the headspace above the gel with a pH 10 buffer 
solution (Figure 3.4). Initially, the entire gel had the bluish color characteristic of 
vesicles. Within 1 h, we could observe that a portion of the gel closest to the buffer 
solution had cleared up ? it was no longer bluish. As time progressed, the clear front 
advanced through the gel and by about 9 h, almost the entire volume of the gel had 
become clear. We also did a control experiment with a pH 8.3 buffer instead of a pH 10 
buffer ? in this case, no perceptible visual changes were observed in the gel over more 
than 48 h. These results imply that in the case of the pH 10 buffer the diffusion of the 
buffer into the gel disrupts the vesicles and transforms them into micelles. Due to the 
weaker light scattering of micelles, the micellized region of the gel shows up as 
practically clear. Incidentally, the photographs in Figure 4 were taken after removing the 
buffer solution from the vial headspace ? this was necessary to clearly visualize the gel in 
the vial. Also, the vial is shown inverted in all the photographs ? this is to indicate that 
the gelatin gel retains its integrity and mechanical strength during the pH-induced 
transition (there is no change in gel volume either).    
 
 
 
 
 49 
 
Figure 3.4.  Movement of micellar front within a vesicle gel due to diffusion of pH 10 
buffer. The buffer is introduced into the headspace above the gel at time zero. As the 
buffer diffuses into the gel, NaOA vesicles in the gel are converted into NaOA micelles, 
as indicated by the change in a portion of the gel from bluish to colorless. As time 
progresses, the micellar front travels deeper into the gel, indicating that more of the 
vesicles are converted into micelles. After 9 h, the micelle region covers most of the gel. 
The plot on the right shows the interface position (measured from the top of the vial) as a 
function of time. The line through the data is a fit to eq 1. 
 
  
 The above result is reproducible and can be replicated in other geometries (see 
below). More quantitative results were gathered from a second experiment in the same 
vial geometry and are shown in the right-hand panel of Figure 3.4. In this case, the 
vesicle-gel was loaded in a cylindrical vial of diameter 7 mm and up to a height of 19 
mm. The remaining volume of the vial was filled with pH 10 buffer. The setup was 
monitored by a camera over the duration of the experiment. From photographs at 
different instants of time, we were able to monitor the movement of the micelle-vesicle 
front (i.e., the interface between the micelle-rich and vesicle-rich regions) within the gel. 
The data (Figure 4) follow the classic diffusive scaling with t .65-67 In other words, the 
 50 
progression of the front is apparently limited by the rate of diffusion of buffer into the 
gel; in comparison, the vesicle-to-micelle transition occurs much faster.  
 
We can then model the time-dependent position of the vesicle-micelle interface 
h(t) by the 1-dimensional diffusion equation:66       
                                               Dthth 2)( 0 ?=                                                    (3.1) 
where D is the diffusivity of the buffer and h0 is the position at t = 0 (which equals the 
total height of the gel). By fitting this equation to the data in Figure 3.4, we obtain a 
diffusivity D of 4.2 ? 10?5 cm2/s for the buffer (hydroxide) ions from the external bath 
into the gelatin gel. This value is quite comparable to the bulk diffusivity of hydroxide 
ions in water (5.6 ? 10?5 cm2/s).68 In other words, the ions diffuse in the gel much like 
they would in water; this is not surprising since the hydrated size of the ions should be 
much smaller than the mesh size of the gel. The latter value is estimated from the 
literature to be about 5 nm for a 5% gelatin gel.69-70   
   
We have seen from Figure 3.3 that it is possible to create a cylindrical gel with 
different microstructures over different regions. For example, half of the cylinder could 
contain vesicles and the other half micelles. If the gel is removed from the buffer solution 
and stored separately, this ?pattern? is retained for several days. Similar patterning can 
also be done with gels of other geometries. For example, we made a vesicle-loaded gel 
(5% gelatin, 0.5% vesicles) into a spherical ball, 41 mm in diameter. This ball was then 
immersed in a bath of pH 10 buffer. As the vesicle-micelle front moved radially inward, 
the vesicles in the outer shell were transformed into micelles whereas the core remained 
 51 
intact. This is evident from Figure 3.5 (top panel) where we see the clear micellar shell 
surrounding the bluish vesicle core. With increasing time, the shell becomes thicker and 
eventually the entire gel contains only micelles. Thus, we can easily create core-shell 
patterns with a vesicle core of given size surrounded by a micellar shell.  
 
Figure 3.5. Patterning of vesicle gels by inducing localized vesicle to micelle transitions. 
(Top panel): core-shell structure with a vesicle-rich core surrounded by a micelle-rich 
shell. This is accomplished by immersing a spherical gel (41 mm diameter) in a pH 10 
buffer. The photographs correspond to increasing incubation times in this buffer solution. 
(Bottom panel): gel in a petri dish with micelle-rich regions created at discrete points. 
This was done by sticking straws into the gel and subsequently filling the straws with the 
pH 10 buffer. The photographs again correspond to increasing times and show the 
expansion of the micelle-rich regions as the buffer diffuses into the gel.    
 
 
Another patterning experiment was carried out with the above vesicle-gel in a 
Petri dish of diameter 90 mm and with the gel thickness being ~ 10 mm. We then stuck 6 
drinking straws loosely into the gel at several locations. The straws were then filled with 
 52 
pH 10 buffer. As the buffer diffused into the gel, the regions surrounding the straws 
became clear, indicating that the vesicles in those regions had been converted into 
micelles. This is shown in Figure 3.5, bottom panel. The clear regions are roughly 
circular in cross-section, and as time progresses, the circles grow radially outward from 
the straw positions. Eventually, the clear regions merge with one another and at this stage 
most of the vesicles in the gel have been transformed into micelles. Variations of the 
above experiments can be used to create more complex patterns. As mentioned in the 
Introduction, an ability to create gels containing pockets of vesicles at precise locations 
within the bulk structure could have applications in areas such as tissue engineering.  
 
One comment needs to be made regarding the reversibility of the transition, i.e., 
back from micelles to vesicles. While a vesicle to micelle transition can be readily 
induced by high pH buffer, the vesicles cannot be subsequently re-formed in the gel if it 
is brought into contact with a low pH buffer. The reason has to do with the mesh size of 
the gel, which was estimated above to be ~ 5 nm. Vesicles of NaOA are thus large 
enough to be trapped within the gel mesh, but spherical micelles of NaOA, which have a 
size around 4-5 nm can ?leak? out of the gel and into the external buffer, thereby 
depleting the NaOA in the interior of the gel. Moreover, for vesicles to be re-formed 
within the gel matrix, several micelles will need to approach and fuse: again, the mesh is 
too dense to facilitate such fusion. To re-form vesicles, one would thus have to melt the 
gel and then combine it with NaOA at the appropriate pH.    
     
 53 
Controlled Release from Vesicle-Gels. Finally, we study the controlled release of a dye 
(calcein) from vesicle-loaded gels and we examine whether the release kinetics are 
influenced by pH. First, Figure 3.6 compares the release of dye from a control gel (no 
vesicles) and from a vesicle-loaded gel, both containing the same amount of dye. In this 
experiment, the pH of both the gel and the external solution is at 8.3. In the case of the 
vesicle-gel, the dye is encapsulated in the vesicles before the vesicles are embedded in 
the gel (i.e., most of the dye is inside the vesicles at time zero). Figure 3.7 shows that the 
vesicle-gel releases dye at a slower rate than the control gel. The slower kinetics are 
evidently due to the additional transport barrier presented by the vesicle bilayers, which 
the dye molecules must first traverse before releasing into the gel and thereafter into the 
external solution. Similar results have been found in other studies.51-56  
 
Figure 3.6. Release profiles of calcein dye from a gelatin gel and from a vesicle-loaded 
gelatin gel. In the case of the vesicle-gel, the dye was encapsulated within the vesicles, 
which were then embedded within the gel. The data show a slower release of dye from 
the vesicle-gel relative to the control gelatin gel. After 10 h, Triton X-100 detergent was 
added to the external solution for both samples (this point is marked by the arrow). The 
detergent diffused in and disrupted the vesicles, which lead to an increase in the rate of 
dye release.  
 54 
Figure 3.6 also shows the effect of Triton X-100 detergent on the release profiles. 
This detergent is known to disrupt vesicle bilayers and thereby convert vesicles into 
micelles.56 We added Triton to the external solution at the 10 h mark. This had no 
significant effect on dye release from the control gel, but the release curve from the 
vesicle-gel shows a sharp increase in slope. Evidently, the Triton molecules diffuse into 
the vesicle-gel and disrupt the vesicles, causing the encapsulated dye to spill out into the 
external gel and thus exit the gel at a faster rate; similar results have been reported 
previously.56 Eventually, the vesicle-gel release profile catches up with that from the 
control gel (4 h after Triton addition). The overlap of the two curves confirms that the 
two gels contain approximately the same overall amount of dye. The Triton experiment is 
also an indirect proof for the existence of intact vesicles in the vesicle-gel. In addition, 
the experiment shows how the release rate from a vesicle-gel can be tuned by addition of 
certain molecules to the external solution.  
   
  We now discuss the effect of external pH on the calcein release profiles. The 
results are shown in Figure 3.7, where data from three experiments are displayed. Two of 
these are identical to those in Figure 3.6, namely a control gel (no vesicles) and a vesicle-
 loaded gel. For these two cases, both the gel and the external solution are at a pH of 8.3. 
Comparing the results, again the vesicle-gel releases dye much more slowly than the 
control gel, consistent with the data in Figure 3.6. The third case is the vesicle-gel placed 
in contact with a pH 10 buffer at time zero: in this case, the release rate is higher 
compared to the same gel at pH 8.3. This result agrees with our expectations and is 
similar to the effect of Triton in Figure 3.6. We expect the pH 10 buffer to diffuse into the 
 55 
vesicle-gel and transform the vesicles into micelles, as shown earlier by the moving front 
in Figure 3.3. In turn, the dye encapsulated in the vesicles will be released into the gel 
matrix and these molecules would no longer have to contend with the vesicle bilayer as a 
transport barrier. This explains the net faster release of dye molecules out of the gel. In 
effect, the dye release kinetics for the pH 10 case gradually begins to resemble that of the 
control gel (i.e., a gel with no vesicles). Figure 3.7 thus conceptually demonstrates the 
use of pH as a trigger to accelerate the release rate of dye from our vesicle-gels. 
 
Figure 3.7.  Tunable calcein release from a vesicle-loaded gel based on the pH of the 
external solution. The control is a gelatin gel surrounded by a pH 8.3 buffer and in this 
case (red circles), the dye is released rapidly. The same gel loaded with vesicles releases 
dye much more gradually at pH 8.3 (green hexagons): in this case, the vesicles are intact 
and the vesicle bilayer thus presents a transport resistance. On the other hand, if the same 
vesicle-loaded gel is placed in contact with pH 10 buffer, the dye release is more rapid 
(blue triangles): in this case, the high pH converts the vesicles into micelles, thereby 
eliminating the transport resistance due to the vesicle bilayers. All the data are fit to eq 2 
and the fit parameters are shown in Table 1.  
 
 
 56 
The dye release curves in Figure 3.7 can be quantitatively treated using the 
following equation due to Peppas:65-67  
                                                nt kt
 M
 M
 =
 ?
                                                   (3.2) 
where Mt /M? is the fractional dye release, k is a rate constant, and n is an exponent 
characteristic of the transport mechanism. This equation is only valid for short times such 
that Mt /M?< 0.6. Fits of eq 2 to the data are shown in Figure 3.7 and the fit parameters 
are shown in Table 3.1 (in all cases, the quality of the fit was very good, with R2 > 0.99). 
The exponent n is determined to be 0.49 for the control gel and 0.53 for the vesicle-gel at 
pH 8.3; both are close to the value of 0.5 expected for Fickian diffusion in a slab 
geometry.66 For the vesicle-gel at pH 10 the value of n is 0.45. The slight discrepancy in 
this case is probably because two processes are occurring simultaneously: the vesicles 
being disrupted into micelles and thus releasing dye into the gel, and the free dye in the 
gel diffusing out into the solution. Note that the rate constant k is highest for the control 
gel and lowest for the vesicle-gel with intact vesicles (pH 8.3). The acceleration of dye 
release due to the increase in pH is reflected in a 50% increase in the magnitude of k 
(from 0.2 at pH 8.3 to 0.3 at pH 10).    
 
Sample k n 
(1) Gel, no vesicles 0.4 0.49 
(2) Vesicle gel, pH 8.3 0.2 0.53 
(3) Vesicle gel, pH 10 0.3 0.45 
 
Table 3.1. Parameters obtained by fitting eq 2 to dye release curves in Figure 7. 
 
 57 
3.4 Conclusions 
We have shown that nanoscale vesicles of NaOA can be entrapped within gelatin 
hydrogels. The resulting vesicle-gel hybrids exhibit the pH-responsive properties of the 
NaOA vesicles. Specifically, when exposed to a pH 10 buffer, the vesicles within the gel 
become transformed into micelles. Vesicle disruption can be done in a controlled manner 
at specific locations within a gel. Gels can thus be ?patterned? to have vesicle-rich and 
micelle-rich domains in predetermined arrangements. The utility of entrapping pH-
 responsive structures within the gel is in the area of controlled release of hydrophilic 
solutes. We show that the release of calcein dye out of a vesicle-gel into the external 
solution is accelerated when the solution pH is raised to 10. This increase is attributed to 
a pH-induced vesicle to micelle transition within the gel, which reduces the transport 
resistance to dye diffusion.    
 
 
 
 
 
 
 
 
 
 
 58 
Chapter 4 
Biopolymer Capsules Containing Vesicles: 
?Motherships? for Controlled Drug Release  
 
 
4.1 Introduction 
 
Two major classes of drug delivery vehicles are (1) microcapsules and (2) 
liposomes. Polymer microcapsules, which can range from a few ?m to >1,000 ?m are 
particularly interesting due to the wide range of packaging, targeting, and tunable release 
mechanisms they offer.71 However, they are often avoided during pharmaceutical 
formulation as many methods of preparing them involve heat or organic solvents, which 
may destroy labile therapeutic molecules.72-73 These common, yet labor-intensive and/or 
drug damaging methods of microcapsule synthesis include layer-by-layer polymer 
deposition (involving burning or harshly dissolving away an inorganic core),74 oil-in-
 water emulsion polymerization,75 and organic solvent evaporation.76 In addition to the 
directly negative potential effects on the drug payload during processing, it can be 
difficult to develop uniformly sized populations of microcapsules, resulting in 
unpredictable drug release and biodistribution profiles.77  
 
Liposomes, the other major class of drug carrier, are generally much smaller in 
size at ~100 nm. These structures have also been suggested for numerous drug delivery 
applications, especially those involving the targeting and uptake of the liposome directly 
into cells.78-79 These self-assembled spheroid structures, with lipid bilayers reminiscent of 
 59 
cell membranes, are excellent for packaging a wide range of therapeutic molecules and 
are highly biocompatible. While PEGylation technology has greatly improved the 
parenteral administration of liposomes,80 these structures still face unfavorable 
destabilization due to absorped proteins from the bloodstream or from roaming 
phospholipases in soft tissue applications. 81 
 
Hybrids of these 2 structures, i.e. capsules containing vesicles, are attractive 
because they may potentially retain the advantages of both, while simultaneously 
mitigating some of the disadvantages of each separately. For example, capsules may 
protect vesicles from external stimuli, such as low or high pH, and from destruction by 
biological ?watchdogs,? e.g. macrophages. Conversely, vesicles may extend the 
functionality of capsules, since they are generally more effective at encapsulating 
hydrophobic drugs (such as many anti-cancer formulations)82-83 than the capsules 
themselves. Finally, the combination of transport resistances from the vesicle bilayer and 
the capsule shell can again prolong the release of encapsulated drug.  
 
In recognition of this potential, such hybrid structures have been synthesized in a 
number of previous studies. The Langer group were the first to show that vesicles could 
be trapped within alginate microcapsules and that dye encapsulated in the vesicles could 
be released slowly under certain conditions.84 Subsequent studies from the same group 
showed that micro-encapsulated vesicles could extend the in vivo drug release time and 
minimize immune responses.85 The concept of ?vesicles in capsules? has been used 
thereafter by other researchers. 86-88 Cohen et al. encapsulated vesicles into alginate-PLL 
 60 
capsules.88 Additionally, Feng et al have demonstrated stable liposome-in-microsphere 
structures which are able to achieve sustained release of drug.89 
 
More recently, Dhoot et al. have studied the effect of crosslinking ions on release 
rates of fluorescein-isothiocyanate labeled bovine serum albumin (FITC-BSA) from 
vesicle-loaded alginate capsules.90 A rapid initial burst of the FITC-BSA protein was 
observed when alginate beads were ionically crosslinked with Ca2+, Al3+, and Ba2+. The 
release rate of the protein was found to differ based on the type of ions used in generating 
the capsules.   Additionally, Ramadas et al. have used vesicle-loaded alginate-chitosan 
capsules for oral delivery of insulin.91 These capsules were able to deliver insulin to the 
intestine without undergoing degradation in the acidic environment of the stomach. This 
example hints at the oral drug delivery administration route as very relevant clinical niche 
for these vesicle-in-capsule formulations. The capsule shell can serve as a protective 
barrier for the vesicles and their encapsulated contents against the harsh gastrointenstinal 
environment while en route to the bloodstream via the intenstine.  
 
In this study, we also create biopolymer capsules containing liposomes, here 
through a mechanism of self-assembly. Liquid mixtures of liposomes and chitosan, a 
cationic biopolymer, are added dropwise into solutions of anionic biopolymer, gellan 
gum, or surfactant, sodium dodecylbenzene sulfonate (SDBS). Spherical capsules are 
formed immediately due to electrostatic interaction between the positively and negatively 
charged structural molecules at the droplet interface. We call these biopolymer capsules 
containing vesicles ?motherships,? as these larger containers are able to retain smaller 
 61 
intact containers, i.e. ?babyships,? within their lumen. An overview of this type of soft 
structure is shown in Figure 4.1. Note that we can create small micro-sized capsules 
(~200 ?m) by means of employing an electric field beneath a 200 ?m nozzle, or we can 
simply drop chitosan/liposome solution out of a standard pipet and create macro-, or 
?over-the-counter? sized capsules. To our knowledge, no one has produced chitosan-
 gellan or chitosan-SDBS capsules containing liposomes. 
 
 
 62 
 
Figure 4.1. Formation Chitosan ?mothership? capsules containing vesicles. Capsules 
are formed by dropping a positively-charged amphiphilic biopolymer mixed with vesicles 
(~100 nm in size) into a solution of negatively charged SDBS surfactant or negatively 
charged gellan gum biopolymer. Spherical capsules are formed instantaneously. Capsule 
size is dictated by droplet size and sizes can range from ~100 ?m (micro) to 10,000 ?m 
(macro) 
 
 63 
Although many advantages of the combined properties of vesicles and capsules 
have already been reported in the literature, here we aim to reiterate some of those 
advantages while extending the potential use of hybrid systems in two key ways. The first 
is that we explore the ability of these motherships to conduct controlled or triggered 
release of the vesicles themselves,92 which is a largely unexplored subtopic within the 
field of drug delivery. Why is controlled release of vesicles potentially high impact from 
a therapeutic standpoint? As a first example scenario outlined in Figure 4.2, these 
capsules could be useful for cancer therapy by injecting or implanting them in the vicinity 
of tumor tissue. An anti-angiogenic drug could be packaged within the lumen of the 
capsule, whereas a cytotoxic cancer drug could be contained within the vesicles. The 
anti-angiogenic drug would be released first and subsequently slow or stop the growth of 
new blood vessels into the tumor tissue.93 Secondly the capsule would degrade, releasing 
the drug-loaded vesicles into the blockaded tumor tissue, locally destroying the diseased 
cells. As a further example, neural circuits are largely dependent upon the triggered 
release of vesicles carrying neurotransmitters within the neuronal synapse.94 Creating 
structures that would allow for triggered release of vesicles may provide a way to build 
high-level circuitry within soft, ?squishy? environments, much like the brain. Of course, 
these sorts of advances may give important insights on neurodegenerative diseases that 
involve hindered neuronal signal transmission (e.g. Multiple Sclerosis, Parkison?s 
Disease).95-96 
 64 
 
Figure 4.2. Schematic of Chitosan ?mothership? capsules attacking a tumor. In step 
(1), the microcapsule ?motherships? would be docked at the tumor site using antibodies 
or magnetic targeting and thereafter release one drug from the capsule lumen (e.g. an 
anti-angiogenic drug). In step (2), capsule shell would become ruptured due to enzymatic 
degradation, and ?babyship? vesicles carrying cytotoxic drug would escape freely into the 
tumor  tissue. 
 
The second extension upon the vesicle-within-capsule concept presented here is a 
vesicle-within-(small)capsule-within-(large)capsule concept. These latter structures are 
built by the exact same one-step drop method, but instead, utilizing nozzles of different 
sizes to put smaller capsules containing vesicles within larger sized capsules, and hence 
provide a simple means of assembling materials display important structural features over 
a hierarchy of length scales. What advantages or insights could these kinds of hierarchical 
structures present? Certainly in biology, there are many examples of systems exhibiting 
important features over a hierarchy of length scales. Cells contain hundreds of smaller 
enclosed structures within their membrane including, the nucleus, endosomal vesicles, 
lysosomal vesicles, mitochondria, etc. Interestingly, an encapsulation system containing 
biological components, such as enzymes, proteins and detoxicants, was originally 
 65 
proposed in 1964 by Chang et al as an ?Artificial Cell.?97 As a next generation of this 
original theory, the hierarchical capsules shown here could potentially serve as much 
more advanced cellular mimics. By the described one-step drop procedure, a pre-
 designed fleet of containers, e.g. one medium-sized capsule containing DNA (nucleus), 
~30 smaller capsules stuffed with ATP (mitochondria), and hundreds of vesicles packed 
with digestive enzymes (lysosomes), could all be housed within one large capsule. Such a 
structure could be prepared in just minutes using only elementary lab hardware. 
 
 
4.2 Experimental Section 
Materials. Acetic acid, sodium dodecyl benzenesulfonate (SDBS), n-dodecyl aldehyde, 
sodium cyanoborohydride, 5,6-Carboxyfluorescein (CF), Triton X-100, rabbit IgG 
(tagged ? = 600 nm emission) and anti-rabbit IgG (tagged ? = 520 nm emission), 
glutaralehyde (40% in water) and chitosanase from streptomyces griseus were all 
purchased from Sigma-Aldrich chemicals. L-?-phosphatidylcholine (PC) was purchased 
from Avanti Polar Lipids, Inc. The magnetic nanoparticles (?-Fe2O3) were purchased 
from Alfa Aesar. Their size was specified to be 32 ? 18 nm, and their average surface 
area was 42 m2/g. Magnets were obtained from United Nuclear. All experiments were 
performed using 18.2 M? DI Water.  
 
Chitosan. Chitosan of medium molecular weight (190-310K) and Brookfield viscosity of 
286 cps was obtained from Aldrich. The reported degree of deacetylation was about 80%, 
and this has been verified by NMR (Supplemental Info). The chitosan backbone is thus 
 66 
mostly composed of D-glucosamine [?-(1,4)-2-deoxy-2-amino-D-glucopyranose] sugars, 
with a small fraction of N-acetyl-D-glucosamine [?-(1,4)-2-deoxy-2-acetamido-D-
 glucopyranose] sugars as well. Chitosan is pH-sensitive as a result of its amine groups 
and is soluble only under acidic conditions, that is, at a pH < 6.5. We have used 0.2M 
acetic acid to control the pH in chitosan solutions.98 Chitosan acts a cationic 
polyelectrolyte under these conditions. 
 
Gellan Gum. Gelrite? Gellan Gum (FW 1,000,000) was obtained from Sigma. Gellan 
gum is a water-soluble negatively charged polysaccharide with a repeating 
tetrasaccharide unit consisting of two residues of D-glucose, one residue of L-rhamnose, 
and one residue of D-glucuronic acid. All units are joined by ?-1,3 glycosidic bonds.  
 
Synthesis of Hydrophobically-Modified Chitosan (hm-Chitosan). We attached n-
 dodecyl tails to chitosan by reacting the amine groups with n-dodecyl aldehyde. The 
procedure follows that reported in the literature.99 Briefly, it involves the addition of 
aldehyde to an acidic solution in a water-ethanol mixture, followed by addition of sodium 
cyanoborohydride. The molar ratio of aldehyde to that of the chitosan monomer(s) was 
fixed at 5% in this study. The hm-chitosan was precipitated by raising the pH and adding 
ethanol, and the precipitate was purified by washing with ethanol followed by deionized 
water. The final hm-chitosan precipitate was redissolved in acetic acid solution, and the 
concentration was recalibrated.  
 
 67 
Preparation of Capsules: In general, capsules were formed by dropping liquid mixtures 
of liposomes and chitosan (?feeders?) into solutions of either negatively biopolymer 
(Gellan Gum) or surfactant (SDBS). The size of the capsule is dictated by droplet size. 
Macro-sized capsules (1000 ? 5000 ?m) were made by dispensing chitosan/liposome 
solution from either a 22G needle via syringe, or by a large aperature fluid transfer pipet. 
Smaller capsules (200, 400 ?m) were made using a Nisco Engineering Encapsulator Unit 
(Norway) with the following settings: voltage 6.5 kV, flow rate 20 mL/hr, 0.2 or 0.4 mm 
(accordingly) needle diameter, needle distance from gellan or SDBS bath 1.3 cm. 
Chitosan-SDBS Capsules. Chitosan (0.5 wt%) was mixed with PC liposomes (0.5 wt %) 
as the feeder. This feeder was loaded into a syringe and dispensed via 22G needle into a 
free-standing solution of 2.5 wt% SDBS. Capsules were allowed to incubate in SDBS 
solution for 2 min, after which the SDBS solution was dumped from the vial while the 
capsules were retained via strainer. Capsules were then washed 3? with 10 ml of DI 
water so as to remove any excess SDBS. In another sample preparation, hm-chitosan (0.5 
wt%) was used to form the capsules instead of unmodified chitosan.  
Chitosan-Gellan Capsules. This method of capsule formation largely follows that of 
Yamamoto el al.100-101 Chitosan (1.5 wt%) was mixed with PC liposomes (0.5 wt%) as 
the feeder. This feeder was also loaded into syringe and dispensed via 22G needle into a 
heated solution of gellan gum (60?C) under stirring. Capsules were allowed to incubate in 
gellan gum solution for 30 min, after which the gellan was dumped from the vial while 
the capsules were retained via strainer. Capsules were then washed 3? with 10 ml of DI 
water so as to remove any excess gellan. After resuspension in buffer solution, the 
capsules were kept under light stirring. 
 68 
 
Lipsome Preparation. Liposomes were prepared by an extrusion method, as 
recommended by the manufacturer (Avanti Polar Lipids). Briefly, dried films of the lipids 
were hydrated under moderate stirring, freeze-thawed 5 times, and then passed through 
two double-stacked polycarbonate membrane filters (100 nm pores) using a Lipex 
pressurized extrusion system.  Dye-filled liposomes were prepared in DI water from PC 
(20 mM) and CF (20 mM) and purified of free CF using the Sephadex G-50 column. 
 
Optical Microscopy. The Zeiss Axiovert 135 TV inverted microscope equipped with the 
Motic Image Plus imaging system has been used for optical microscopy (bright-field). 
Capsules were imaged with a 2.5X objective.  
 
Dynamic Light Scattering (DLS). DLS was used to characterize the sizes of vesicles in 
solution. A Photocor-FC light scattering instrument with a 5 mW laser source at 633 nm 
was used at a scattering angle of 90?. A logarithmic correlator was used to measure the 
intensity autocorrelation function. The hydrodynamic size of the vesicles was extracted 
from the data using the Stokes-Einstein equation.   
 
Fluorescence Microscopy. Photomicrographs of fluorescent electrodeposited materials 
were taken by a fluorescence stereomicroscope (MZFLIII, Leica) equipped with a digital 
camera (Spot 32, Diagnostic Instruments). To observe primary IgG antibody fluorescence, 
the filters were chosen with an excitation wavelength of 560 nm (bandwidth of 40 nm), 
and an emission filter of 610 nm. To observe secondary anit-IgG antibody fluorescence, 
 69 
the microscope was set with an excitation wavelength of 480 nm (bandwidth of 40 nm) 
and a long-pass emission filter at 510 nm.  
 
Release Profiles. Lipid vesicles were combined with 25 mM CF dye. To separate 
unencapsulated dye, the vesicle-dye mixture was passed through a Sephadex G-50 size-
 exclusion chromatography (SEC) column. The vesicle fraction (with encapsulated dye) 
was collected and used in preparing the vesicle-loaded capsules. Note that in these 
experiments, ?macrocapsules? approximately 5 mm in diameter were used so as to allow 
easy collection of external aqueous media. To examine the release of dye, 4 g of 
phosphate buffer solution (pH 7.4) was added to the capsules after washing. The 
concentration of CF in the external buffer solution was measured by UV-Vis 
spectrometry (Cary Bio 50). A control gel (no vesicles) was prepared using an amount of 
CF equal to that in the encapsulated vesicles. To determine the latter, 2 mL of the vesicle 
fraction from SEC was analyzed by UV-Vis after the vesicles were disrupted by adding 
50 ?L of Triton-X100.    
 
Enzymatic Degradation of Capsules. Chitosanase was added to a Petri dish containing 
several chitosan-gellan capsules at a concentration of 0.125 units/ml in acetate buffer 
solution (pH 5). Capsules were then observed under phase contrast microscopy for up to 
5 days. Control capsules in acetate buffer solution (pH 5) with no enzyme added were 
also observed. 
 
 
 70 
Targetability of Capsules 
Magnetic Capsules. Chitosan (0.5 wt %) and liposomes (0.5 wt%) were mixed with 0.1% 
wt ?-Fe3O4 nanoparticles and subsequently dropped into anionic surfactant or biopolymer 
solution. Capsules were then moved within stagnant fluid in a vial using a large magnet, 
and then against a flowing aqueous stream within Silicone tubing connected to a pump.   
Antibody Conjugation. Primary IgG antibody was added in excess to chitosan-SDBS 
capsules in phosphate buffer solution (pH 7.4) until mild stirring. 100 ?l of 
glutaraldehyde solution was then added to the buffer media under increased stirring. 
Reaction was allowed to take place for 20 min. The buffer solution was then drained from 
the vial according to the straining procedure described above. Capsules were washed 3? 
with PBS and then resuspended in PBS. Fluorescence microscopy was used to detect the 
presence of antibody on the capsule surface. IgG-capsules were then incubated with a 
secondary antibody, anit-IgG, for a period of 30 min. Capsules again washed according to 
the above procedure, and imaged in with Fluorscence microscopy. 
 
4.3 Results and Discussion 
In Figure 4.3, release profiles of model drug, carboxyfluorescein (CF), from these 
mothership structures are shown. Figure 4.3a shows that chitosan-gellan motherships 
impregnated with drug-loaded liposomes (red circles) display a sustained release of drug. 
In contrast, chitosan-gellan control capsules with the same amount of drug freely 
dispersed inside the capsule lumen shows a classic ?dosage dumping? release profile 
(blue circles).  This result is explainable by the additional transport resistance provided 
by the liposome bilayer in the motherships. While drug inside the control capsules must 
 71 
only cross 1 barrier, i.e. the capsule shell, the dye inside the motherships must cross two: 
(1) lipid bilayer and (2) capsule shell. At 3.25 h, a detergent molecule Triton X-100 was 
added to the external buffer and a spike in dye release for mothership capsules is 
observed for the remainder of the 4 timeframe. This result gives two key insights into the 
experiment. Firstly, it demonstrates that the liposomes existed intact inside the 
motherships. Triton X-100 is molecule which rapidly disrupts intact vesicles into mixed 
micelles, henceforth spilling the contents of the vesicle core into external solution. Thus, 
the spike in release implies that intact vesicles were broken and the newly escaped drug 
molecules take on a ?dosage dumping? release profile similar to that observed for the 
control capsules. Secondly, the experiment demonstrates that adequate control capsules 
which contain an approximately equal amount of dye to the mothership capsules were 
properly produced. 
 72 
 
Figure 4.3. Controlled Release of Dye from ?Mothership? Capsules. In (a), a sustained 
release is observed for chitosan-gellan capsules impregnated with carboxyfluorescein-
 loaded liposomes (red circles). Chitosan-gellan control capsules with dye freely dispersed 
inside the capsule lumen shows a ?dosage dumping? release profile (blue circles). 
Addition of Triton X-100 at 3.25 h shows a spike in dye release for ?mothership? 
capsules, indicating vesicle intactness. In (b), chitosan-SDBS capsules impregnated with 
CF-loaded liposomes show (blue circles) show a similar release profile to control 
capsules containing no liposomes (yellow triangles), suggesting potential liposome 
rupture during capsule formation. In contrast, hm-chitosan capsules formed in SDBS 
while retaining liposomes, shows a sustained release over a 2 hour interval. 
 
 
 73 
Figure 4.3b shows the release profile of CF from motherships constructed with 
SDBS surfactant instead of anionic biopolymer. Unexpectedly, the chitosan-SDBS 
motherships impregnated with CF-loaded liposomes (blue circles) show a similar release 
profile to control capsules containing no liposomes (yellow triangles). This result 
suggests potential liposome rupture during capsule formation. Such an outcome is 
feasible considering high enough concentrations of SDBS molecules are able to rupture 
liposomes into mixed micelles. Previous work on a similar polyelectrolyte-surfactant 
system describes the resulting structures as ?gel beads,? and suggests that the surfactant 
quickly diffuses into the entire polyelectrolyte droplet.102-103 While our own optical 
microscopy of these capsules displays a distinct ?shell? structure of the capsule distinct 
from the lumen, it is probable that a large number of surfactant molecules infiltrate the 
droplet core prior to rigid shell formation. In contrast, relative to the unmodified 
chitosan-SDBS motherships, hydrophobically modified (hm)-chitosan motherships 
formed in SDBS while retaining liposomes show a sustained release over a 2 hour 
interval. This could potentially be explained by a delayed diffusion of SDBS into the 
capsule core due to association with the hydrophobic grafts along the chitosan backbone. 
This experiment gives some interesting insight into the process of formation of these 
polyelectrolyte-surfactant complexes, although it also suggests that they are unsuitable 
carriers of lipid vesicles. 
 
 
Figures 4.4 and 4.5 outline the potential targetability of these structures via 
magnetic nanoparticles and antibodies, respectively. In Figure 4.4, magnetically targeted 
 74 
?mothership? capsules via encapsulated ?-Fe3O4 nanoparticles show their magnetic 
properties in response to a bar magnet. Figure 4.4a displays a magnet collecting a pile of 
capsules onto the side of the vial. Indeed, this magnet can be used to rapidly move the 
capsules to any location within the stagnant fluid. Furthermore, as shown in Figure 4.4b, 
these magnetically-loaded motherships are able to be pulled against a fluid flow within a 
tube using the magnet. This level of crude targetability is an efficient payoff relative to 
the simple preparatory work of mixing in the nanoparticles along with the liposomes in 
the feeder solution prior to capsule formation.  
 
 
 
Figure 4.4. Magnetically targeted ?mothership? capsules via encapsulated ferrite 
nanoparticles show their magnetic properties in response to a bar magnet. (a) magnet 
placed near capsules in a vial (b) magnet placed next to a tube in which the capsules are 
flowing along with the fluid (water) from left to right. 
 
 
 75 
 
Figure 4.5. Antibody targeted ?Motherships. (a) Fluorescently tagged primary IgG 
antibody (emission 600 nm) is covalently conjugated to the surface of a Chitosan-SDBS 
?mothership? capsule as shown by fluorescence micrograph. After incubation with a 
secondary anti-IgG antibody and subsequent rinsing, fluorescent signal at 520  nm is 
observed, indicating that a significant amount of primary surface IgG is actively available 
for ?lock-and-key?binding. 
 
Figure 4.5 demonstrates a targetability proof-of-concept via the attachment of 
fluorescently-labeled mouse IgG antibodies onto the surface of mothership capsules. The 
synthesis here exploits the plentiful surface amine groups provide by the chitosan 
molecules. Glutaraldehyde is used as a bifunctional linker, which non-specifically 
crosslinkes moieties bearing amine groups. By adding in glutaraldehyde to a stirring 
mixture of antibody and capsules, many antibodies become attached to the capsule 
surface. This linkage is confirmed via fluorescence microscopy (Figure 4.5a) as a 
uniformly bright red color is observed on the capsule after thorough washing. 
 
To demonstrate functionality of the newly conjugated antibodies, capsules were 
brought incubated for 20 min with a secondary antibody specific for IgG (FL-anti-mouse-
 76 
IgG). After washing several times with buffer, the capsules were examined under 
fluorescence microscopy as shown in Figure 4.5b. After incubation with a secondary anti-
 IgG antibody and subsequent rinsing, fluorescent signal at 520  nm is observed, 
indicating that a significant amount of primary surface IgG is actively available for ?lock-
 and-key? binding, although not with the same level of uniformity observed for the 
primary antibody. This result suggests that a significant amount of the primary antibody 
is available for binding and hence applicable for targeting applications. Of course, a 
remaining portion of the primary antibody is not available for binding, which is not 
surprising given the crude, non-specific nature of glutaraldehyde chemistry involved.  
.  
Next, we explore the triggered rupture of these motherships via enzymatic degradation. 
Figure 4.6a displays a phase micrograph of an intact chitosan-gellan mothership, 
approximately 5 mm, in size prior to addition of chitosanase. After enzyme is added to 
the surrounding media, the progression of mothership degradation over time is shown in 
Figure 4.6b. At 1 hour, little effect is shown on the capsule shell, i.e. the effect of the 
enzyme is not immediate. However, after a period of 2 days the capsule shell has clearly 
shriveled, presumably due to loss of mechanical integrity. Furthermore, after a period of 
5 days, the capsules are completely broken and pieces of shell debris are observed 
floating in the sample. After 7 days, the debris is fully dissolved and clear solution 
produced. It is important to note that after this 7 day period, the solution scatters light 
from structures measured at approximately ~100 nm in size by dynamic light scattering 
(DLS). These structures are presumably intact vesicles, as they closely match the DLS 
measurements of the same vesicles prior to addition to the feeder solution. Control 
 77 
capsules to which no enzyme was added show no degradation after the same time period 
of 5 days. Also, it is worth noting that the chitosanase cannot degrade chitosan capsules 
formed with SDBS, most likely due to protein denaturing caused by the surfactant.  
 
Figure 4.6. Enzymatic Degradation of ?Mothership? Capsules. (a) Phase contrast 
micrograph of Chitosan ? Gellan capsules formed by standard drop method via 22G 
syringe needle; (b) 1h, 2d, and 5d micrographs of chitosan-gellan capsule after addition 
of 10 units/ml of chitosanase in external buffer solution. Capsule appears almost wholly 
degraded after the studied 5 day interval. Note that the chitosanase cannot degrade 
chitosan capsules formed with SDBS. 
 
Lastly, we were interested in building hierarchical structures by placing smaller 
capsules within larger capsules, again employing the same one-step drop method. A 
schematic of this kind of structure is shown in Figure 4.7. By a simple one-step self-
 assembly process, structures displaying a hierarchy of length scales, from nano (i.e. 
vesicles), to micro (microcapsules), to macro (large capsules) can be produced. These 
structures have the capability to package a wide variety of molecules and targeting agents 
either on their surface, or within their multi-level lumens. Presumably the release of each 
of the molecule would differ based upon their original location within the hierarchical 
structure. 
 
 
 78 
 
Figure 4.7. Schematic of hierachical nano-micro-macro ?motherships.? By a simple 
one-step self-assembly process, structures displaying a hierchary of length scales, from 
nano (i.e. vesicles), to micro (microcapsules), to macro (large capsules) can be produced. 
These structures can package a wide variety of molecules and targeting agents either on 
their surface, or within their multi-level lumens. 
 
In Figure 4.8, small chitosan SDBS capsules were formed by the standard drop 
method via 22G syringe needle. After incubation, capsules were removed from SDBS 
solution and placed into a fresh chitosan solution. Chitosan solution containing capsules 
was uptaken by a wide-aperture fluid transfer pipet as a new feeder and dispensed 
dropwise into a solution of SDBS, thus forming capsule-within-capsule structures. As 
shown in the micrographs, various numbers of smaller capsules can be placed within 
these larger capsules. Note that we can create the same sorts of capsule-within-capsule 
structures with chitosan-gellan, although the inner capsules tend to lose some shell 
integrity due to pressure from the chitosan in the capsule lumen. Of course, the advantage 
with the chitosan-gellan system is that we can successfully place intact nanostructures, 
i.e. vesicles, within the inner capsule lumens.  
 
 79 
 
 
Figure 4.8. Phase Microscopy (2.5?) of Capsules-within-capsules. Small Chitosan 
SDBS capsules were formed by the standard drop method via 22G syringe needle. After 
incubation, capsules were removed from SDBS solution and placed into a fresh chitosan 
solution. Chitosan solution containing capsules was uptaken by a wide-aperture fluid 
transfer pipet and dispensed dropwise into a solution of SDBS, thus forming capsule-
 within-capsule structures. 
 
Lastly in Figure 4.9, we study the release of dye from capsules-within-capsules 
using the chitosan-SDBS system. Release of CF dye from smaller capsules packaged 
with larger capsules (red circles) is significantly sustained over a 30 period relative to the 
same number of small capsules freely dispersed in aqueous buffer (blue circles). This is 
expected since the dye molecules experience an added transport barrier of the large 
capsule shell. Similar results are observed for chitosan-gellan capsules-within-capsules 
confirming a proof of concept for controlled release from these hierarchical structures.  
 
 
 80 
 
 
Figure 4.9. Release of dye from capsules-within-capsules. Release of 
carboxyfluorescein dye from smaller capsules packaged with larger capsules (red circles) 
is significantly sustained over a 24 period relative to the same number of small capsules 
freely dispersed in aqueous buffer (blue circles). 
 
4.4 Conclusions 
We have shown that ?mothership? capsules with hierarchical structural features can be 
formed by adding droplets of a feeder solution consisting of liposomes and chitosan, a 
cationic biopolymer, into a solution of anionic biopolymer (gellan gum) or surfactant 
(SDBS). The resulting structures are spherical capsules; assembly is achieved via 
electrostatic interactions between oppositely charged polymers/surfactants at the interface 
of the droplet. Capsule size is simply dictated by drop size. Chitosan-gellan motherships 
are able to sustain model drug release due via intact liposomes carrying the drug within 
the capsule. In contrast, liposomes within chitosan-SDBS motherships disrupt quickly; 
this disruption process can be slowed by the covalent addition of hydrophobes to the 
chitosan backbone prior to capsule formation. The potential for targetability of 
 81 
motherships was confirmed by (1) by dispersing ?-Fe3O4 nanoparticles within the capsule 
lumen and (2) attaching IgG antibodies to the capsule surface, a significant portion of 
which are available for binding secondary antibody. Chitosan-gellan motherships can be 
degraded by the addition of the enzyme chitosanase; complete degradation occurs over a 
7 period, with intact vesicles present in the remaining solution post-degradation. Finally, 
hierarchical capsule-within-capsule structures can be produced with either chitosan-
 SDBS or chitosan-gellan systems. Dye release from smaller capsules entrapped within 
larger capsules undergo sustained drug of a 30 h period.  
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 82 
Chapter 5 
Vesicle Capture on Patterned Surfaces Coated 
with Amphiphilic Biopolymers  
 
 
5.1 Introduction 
 
Vesicles (or liposomes) are self-assembled ?nano-containers? formed by lipids or 
surfactants in aqueous solution.104 These structures are ~ 100 nm in size and comprise an 
aqueous core and a lipid bilayer. The aqueous core can be used to encapsulate 
hydrophilic molecules such as drugs, proteins, or genes, while hydrophobic and 
amphiphilic substances can be integrated into vesicle bilayers.104 Few, if any, other 
nanostructures demonstrate this level of versatility as carriers of useful payloads. 
Accordingly, vesicles have been explored and exploited for a myriad of applications, 
including targeted drug delivery, gene transfection, imaging agents, biosensors, food 
science, and cosmetics.105  
 
Recently, there has been new-found interest into the capture of vesicles on solid 
substrates, as opposed to vesicles in solution.106 The motivation for such studies includes: 
(a) fundamental aspects, e.g., related to vesicles as biological models for adherent 
cells;107-109 as well as (b) applied aspects related to the fabrication of biosensors or 
modified biomaterials.110-113 In this context, it is to be noted that vesicles usually fuse or 
rupture into flat bilayers upon approaching a solid surface.114 This highlights the inherent 
disconnect between the ?soft? nature of vesicles and the ?hard? nature of solid surfaces. 
 83 
However, it is often desirable to capture intact vesicles on surfaces.115-121 If vesicles were 
intact, their internal volume would be available for entrapping biomolecules, drugs, or 
fluorescent molecules, which could be useful for sensor and immunoassay applications. 
In addition, proteins embedded in vesicle bilayers are expected to more closely mimic 
their in vivo function compared to the same proteins in supported planar bilayers.116,122  
 
Previous attempts to capture intact vesicles on surfaces have employed DNA 
tethering,115-117 covalent binding to gold or polystyrene,118-119 or biotin-streptavidin 
linking schemes.120-121 These methods generally involve labor-intensive experimental 
procedures and expensive reagents. Furthermore, spatial and temporal control of vesicle 
capture (i.e., onto specific areas of a given surface at a given time) cannot be easily 
achieved by these methods. A greater level of control could be advantageous for many 
applications ? for example, vesicle capture at predetermined locations (e.g., ?vesicle 
arrays?) could facilitate the development of new biosensors.116    
 
In this study, we explore a simple and inexpensive method to capture vesicles 
onto solid surfaces with a high degree of spatio-temporal control. Our method exploits 
two sets of key results from our laboratories over the past several years. The first is the 
ability to electrodeposit the biopolymer chitosan onto patterned surfaces using electrical 
signals.123-126 Chitosan is a widely available cationic biopolymer that has a pH-responsive 
character (pKa ~ 6.0) ? i.e., it transforms from a soluble to an insoluble form upon 
increasing pH above ca. 10. Accordingly, chitosan films can be deposited on cathode 
surfaces upon application of a current due to the high local pH near this electrode.123-125  
 84 
Here, we extend the electrodeposition scheme to a derivative of chitosan that is 
functionalized with hydrophobic tails (Figure 5.1). We term this derivative as 
?hydrophobically modified chitosan?, or hm-chitosan for short.99 Previously, we have 
studied the interaction between hm-chitosan and vesicles in solution.99,127-128 The key 
result was that the hydrophobes from this amphiphilic biopolymer tended to insert into 
the hydrophobic bilayers of vesicles, thus transforming the vesicle solution into a ?vesicle 
gel?, with the vesicles now serving as junction points in a network. Here, we show that 
the affinity for vesicles also extends to hm-chitosan films. When a surface coated with 
hm-chitosan is exposed to a solution of vesicles, the film captures vesicles simply via non-
 covalent interactions between the vesicles and the polymer hydrophobes (see schematic 
in Figure 5.1). Thus, hm-chitosan can act as a ?soft? interconnect between vesicles and a 
hard surface such as a gold electrode.126 Moreover, our ability to selectively deposit hm-
 chitosan in predesigned patterns, as shown in Figure 5.1, allows us to create 
corresponding patterns of immobilized vesicles.   
 
 
 
 85 
 
Figure 5.1. Schematic showing the spatiotemporal capture of vesicles using hm-chitosan. 
The structure of hm-chitosan as well as a schematic of its chain are shown in the top left. 
The chain has a hydrophilic backbone shown in blue and pendant hydrophobes (C12 tails) 
that are shown in red. In step (1), thin films of hm-chitosan are electrodeposited onto gold 
cathodes that are patterned on a silicon chip. Next, in step (2), the chip is incubated with a 
solution of vesicles. Intact vesicles become spontaneously anchored onto the polymer 
films. The inset shows the likely mechanism for such anchoring, which is via non-
 covalent interaction between the hydrophobes on the polymer and vesicle bilayers.  
 
 
It is worth emphasizing certain features of our approach as well as some potential 
applications. Our approach (Figure 5.1) involves deposition of hm-chitosan followed by 
capture of vesicles on the surfaces of these films. The advantage is that the vesicles 
remain accessible to the external environment, which facilitates their use as sensors. (The 
accessibility of vesicles is more problematic if the vesicles are simply co-deposited with 
the polymer, as has been done earlier.129) We will also present several pieces of evidence 
to show that the vesicles are indeed intact, i.e., they are not being ruptured into flat lipid 
bilayers within the hm-chitosan film. Thus, the payload within the vesicles is expected to 
remain available for future use. The latter aspect is important in considering potential 
applications for vesicle-polymer hybrid films in drug delivery or wound healing. For 
 86 
example, in the case of chronic-wound healing, macroscopic films of hm-chitosan could 
be therapeutically functionalized with vesicles housing several types of growth factors 
(i.e., proteins that promote wound healing). These films could then be used to cover 
chronic wounds while protecting them from exudation or infection (chitosan is known 
also for its anti-bacterial properties). In turn, the controlled release of the growth factors 
from the vesicles attached to the films would enhance and sustain the wound-healing 
process. Note also that the bioactivity of growth factors is likely to be maintained when 
they are within vesicles; in contrast, growth factors embedded directly within polymer 
films may lose some of their activity.130 In sum, vesicle-polymer hybrids can have 
interesting applications, and the present study shows how these materials can be readily 
assembled via non-covalent hydrophobic interactions.  
 
 
5.2 Experimental Section 
Materials. Chitosan of medium molecular weight (190-310K) and Brookfield viscosity 
of 286 cps was obtained from Sigma-Aldrich. The reported degree of deacetylation was 
about 80%. Chitosan is soluble only under acidic conditions (pH < 6.5) and here it was 
dissolved in 0.2 M acetic acid. The phoshpolipids L-?-phosphatidylcholine (PC) and 
biotinylated phosphatidylethanolamine (PE-biot), and the fluorescent lipid 1'-dioctadecyl-
 3,3,3',3'-tetramethylindocarbocyanine perchlorate (DiI) were purchased from Avanti 
Polar Lipids. The surfactants cetyltrimethylammonium tosylate (CTAT), sodium dodecyl 
benzenesulfonate (SDBS), and Triton X-100, the dye 5,6-carboxyfluorescein (CF), and 
the reagent n-dodecyl aldehyde were purchased from Sigma-Aldrich. Streptavidin bound 
 87 
to fluorescein isothiocyanate (FITC-streptavidin) and the succinimidyl ester of CF (NHS-
 fluorescein) were purchased from Fluka. All experiments were performed using distilled-
 deionized (DI) water.  
 
Synthesis of Chitosan Derivatives. The hm-chitosan was synthesized by attaching 
n-dodecyl tails to the chitosan backbone via reaction with n-dodecyl aldehyde. The 
procedure has been reported in our earlier paper99 and follows that described in the 
literature.17,98,131 The degree of hydrophobic substitution follows the reaction 
stoichiometry and in this study it was fixed at ca. 2.5 mol% of the available amine groups. 
Fluorescently-labeled chitosan and hm-chitosan were synthesized by reacting the 
polymers with NHS-fluorescein, as previously reported in the literature.124  
 
Vesicle and Liposome Preparation. Both surfactant vesicles and lipid vesicles 
(liposomes) have been used in this study. Catanionic surfactant vesicles12,132 were 
prepared by mixing 0.7 wt% of the cationic surfactant CTAT and 0.3 wt% of anionic 
surfactant SDBS (~ 2:1 molar ratio) in DI water and gently stirring overnight. Dye-filled 
catanionic vesicles were prepared by combining 1 mM of CF with the CTAT/SDBS 
mixture, followed by separation of vesicles from free dye using a Sephadex G-50 column 
(from Roche). Liposomes were prepared by an extrusion method, as recommended by the 
manufacturer (Avanti Polar Lipids). Briefly, dried films of the lipids were hydrated under 
moderate stirring, freeze-thawed 5 times, and then passed through two double-stacked 
polycarbonate membrane filters (100 nm pores) using a Lipex pressurized extrusion 
system.  Dye-filled liposomes were prepared in DI water from PC (20 mM) and CF (15 
 88 
mM) and purified of free CF using the Sephadex G-50 column. Fluorescently labeled 
liposomes were prepared in DI water by combining the lipid PC (13 mM) with trace 
amounts (13 ?M) of the fluorescent lipid DiI. For the streptavidin binding assay, 
biotinylated liposomes were prepared by combining PC and PE-Biotin in a molar ratio of 
9:1, with a total lipid concentration of 1 wt%. Vesicle sizes in all cases were measured 
using a Photocor-FC dynamic light scattering (DLS) instrument.   
 
Preparation of Giant Unilamellar Vesicles (GUVs). The GUVs were prepared by 
electroformation, as described in the literature.133-134 The lipids (1 mg/mL) and DiI (7.7 
?g/ml) were dissolved in chloroform. One drop of this solution (5 ?L) was deposited 
onto the conducting side of an indium tin-oxide (ITO)-coated glass slide. The solvent was 
removed first under desiccation for 1 h, and then freeze-drying for 3 h. A chamber was 
then made by creating an O-ring out of Seal-Ease and then pressing a second ITO-coated 
slide, conducting side facing downward, above the original slide. The chamber was 
hydrated with a solution of 100 mM sucrose in DI water via injection needle through the 
Seal-Ease; after injection, the needle was removed and the hole closed by Seal-Ease. 
Alligator clips were connected to both glass slides as well as a function generator (BK 
Precision 10 MHz Sweep/Function Generator 4017) via a BNC connector. An electric 
field of AC 1.5 V at 10 Hz was applied for 2 h at 55?C; the frequency was then dropped 
to 1 Hz for an additional 50 min. The fluid in the chamber now consisted of GUVs. 
Electrodeposition. Electrodeposition was performed on ?chips? fabricated from silicon 
wafers with deposited micropatterns of gold (Figure 5.2). Fabrication procedures have 
been described in detail in our earlier papers.123-125 Electrodeposition was performed by 
 89 
negatively biasing a specific lead on the chip while it was partially immersed in an 
aqueous solution containing 1 wt% of either chitosan or hm-chitosan at a pH ~ 5. A DC 
power supply (model 6641C, Agilent Technologies) was used to supply a constant 
current to the chip and counter electrode over a 2 min period. Photomicrographs of 
fluorescent electrodeposited materials were taken by a fluorescence stereomicroscope 
(MZFLIII, Leica) equipped with a digital camera (Spot 32, Diagnostic Instruments). To 
observe CF or FITC fluorescence, the microscope was set with an excitation wavelength 
of 480 nm (bandwidth of 40 nm) and a long-pass emission filter at 510 nm. To observe 
DiI fluorescence, the filters were chosen with an excitation wavelength of 560 nm 
(bandwidth of 40 nm), and an emission filter of 610 nm. In all cases, Image J1.34S 
software from NIH was used to analyze images and quantify fluorescence intensity.  
 
Cryo-TEM. C-FLAT holey carbon grids with a hole size of 1.2 ?m were purchased from 
Electron Microscopy Sciences. Grids bearing hm-chitosan and surfactant vesicles were 
plunged into liquid ethane (?183?C) using a Gatan CryoPlunge3, so as to form vitrified 
specimens and thereby preserve any molecular assemblies present. The samples were 
thereafter imaged on a JEOL-2100 LaB6 TEM at liquid nitrogen temperature. 
 
 
 
 
 
 
 90 
5.3 Results and Discussion 
Vesicle Capture on hm-chitosan Films. The first step in our approach involves 
electrodeposition of hm-chitosan films onto gold electrodes. These films are then to be 
used to capture vesicles. In the context of vesicle capture, it is useful to compare 
hm-chitosan with the native chitosan (no hydrophobes). Electrodeposition of native 
chitosan was first demonstrated in our laboratories in 2002;123 the present study is the 
first to extend this capability to hm-chitosan. For our initial studies, we used the two-
 electrode chip shown in Figure 5.2. As mentioned earlier, the chip substrate is a silicon 
wafer, on which gold electrodes are deposited in a specific pattern. We then conducted 
sequential electrodeposition of hm-chitosan and chitosan (both tagged with the green 
fluorescent dye, NHS-fluorescein) on the left and right electrodes, respectively. This was 
done as follows: first, the right electrode was connected to the power supply and the other 
unconnected while the chip was immersed in a solution of chitosan. The connections 
were then switched (only left electrode connected) and the chip was immersed in a 
solution of hm-chitosan. After deposition, the electrodes were disconnected from the 
power supply, and the chips were rinsed several times with DI water. Figure 5.3 (top) 
shows the results of the experiment: the green color on both electrodes in the two 
fluorescence images indicates successful deposition of both hm-chitosan and chitosan.  
 
 
 
 
 
 91 
 
Figure 5.2. Photograph of micropatterned chips used in this study. The leads and 
electrodes are formed by depositing gold on a silicon wafer using lithographic techniques. 
On the left is a 2-electrode chip and on the right a 6-electrode chip. A US penny is shown 
for size comparison.  
 
Next we examined the relative capabilities of hm-chitosan and chitosan to capture 
vesicles from solution. These experiments were conducted with both surfactant vesicles 
as well as lipid vesicles (liposomes). The catanionic surfactant vesicles (70/30 
CTAT/SDBS, total 1 wt%) had an average diameter around 120 nm, as measured by DLS. 
The liposomes of PC were made by extrusion through 100 nm membrane filters and had 
an average diameter around 110 nm from DLS. Both the vesicles and the liposomes were 
tagged with the fluorescent lipid DiI, which incorporates into the bilayer membranes of 
the above structures. Note that DiI exhibits a red fluorescence, i.e., a distinct color 
compared to the green signal from the chitosan and hm-chitosan. We incubated the chip 
on the left with surfactant vesicles and the one on the right with the liposomes, both for 
10 min. The chips were then rinsed three times with DI water to remove weakly adsorbed 
structures and then imaged under the fluorescence microscope using red filters. The 
 92 
resulting images (Figure 5.3, bottom) show a significantly higher red fluorescent signal 
from the hm-chitosan side compared to the chitosan-side ? and this is the case for both 
the surfactant vesicles and the liposomes. These results show that hm-chitosan is 
considerably more effective at capturing vesicles than chitosan. We believe this is 
because hydrophobes from hm-chitosan chains can get inserted into vesicle bilayers 
through non-covalent hydrophobic interactions (as depicted in Figure 5.1). Such 
interactions enable a tight anchoring of the vesicles to the polymer film.  
 
 
 
 93 
 
Figure 5.3: Comparison of hm-chitosan (H) and chitosan (C) films in terms of their 
ability to capture vesicles. Top images: Green-filtered fluorescence micrographs showing 
deposition of NHS-tagged hm-chitosan on the left electrode and NHS-tagged chitosan on 
the right electrode of a two-electrode chip. The chips are then incubated with vesicles for 
10 min, followed by rinsing with DI water. Bottom images: Red-filtered fluorescence 
micrographs showing the presence of DiI-tagged vesicles on the electrodes. The results in 
(a) (left-column) are obtained with catanionic surfactant vesicles, while those in (b) 
(right-column) are obtained with liposomes. In both cases, the vesicles are anchored 
preferentially to the hm-chitosan film than to the chitosan one.  
 94 
The superior effectiveness of hm-chitosan in capturing vesicles is a robust result and this 
is further proven by experiments with varying vesicle incubation times. For these 
experiments, surfactant vesicles were used and the time of chip incubation in the vesicle 
solution was varied from 5 to 10 to 20 min. All other experimental variables, including 
rinsing times and image exposure time were kept constant. The results in Figure 5.4 show 
an increase in red fluorescence from the hm-chitosan electrode with increasing incubation 
time ? the increase is quite significant between 5 and 10 min and slight between 10 and 
20 min. In all cases, the signal from the hm-chitosan side dwarfs that from the chitosan 
side. Similar results were obtained for macroscopic films of chitosan and hm-chitosan 
simply dried onto glass slides and incubated with tagged surfactant vesicles for various 
time intervals (data not shown). These results suggest that vesicle capture on hm-chitosan 
proceeds by surface diffusion of vesicles until they connect with free hydrophobes from 
the polymer, whereupon the vesicles get strongly bound. As the free hydrophobes get 
used up, fewer vesicles are able to bind strongly and the binding capacity of the film 
becomes saturated. 
 
 
 95 
 
Figure 5.4: Variation in the extent of vesicle capture based on the time of chip incubation 
with a vesicle solution. Hm-chitosan (H) and chitosan (C) are deposited on the left and 
right electrodes, respectively. The chip is incubated with a solution of surfactant vesicles 
for different lengths of time and then imaged by fluorescence microscopy. The images 
are red-filtered fluorescence micrographs showing DiI-tagged vesicles on the electrodes. 
A constant exposure time of 20 s at the excitation wavelength was used to obtain each 
image. The results show increasing capture of vesicles on the H side and negligible 
capture on the C side. 
 
 
Intactness of Captured Vesicles. Next we tackle the question of whether the vesicles 
captured on hm-chitosan films are intact or whether the red fluorescent signal reflects 
fragments of vesicle bilayers. Figure 5.5 is a first piece of evidence to demonstrate 
intactness of vesicles. In this case, we prepared a solution of fluorescently tagged giant 
unilamellar vesicles (GUVs) of phospholipids. Optical microscopy (not shown) revealed 
that the GUVs ranged in size from 10 to 50 ?m, which is typical of GUVs made via 
electroformation. We then incubated a chip bearing hm-chitosan in the GUV solution for 
10 min, rinsed the chip and observed it under a fluorescence microscope. Note the red 
fluorescent ?hot spots? on the hm-chitosan film ? these are shown at two different levels 
 96 
of magnification in Figures 5.5. The spot sizes compare well with the sizes of GUVs and 
therefore it is likely that the spots correspond to intact GUVs. Thus, large microsized 
GUVs are shown to remain intact when immobilized on an hm-chitosan film.  
 
 
Figure 5.5: Immobilization of giant unilamellar vesicles (GUVs) on an hm-chitosan film. 
After 10 min incubation with DiI-tagged GUVs, multiple red-fluorescent ?hot spots? 
appear on the electrode bearing hm-chitosan. A zoomed-in image of a ?hot-spot? shows 
an intact GUV of ca. 20 ?m diameter. 
 
Returning to the nanosized vesicles and liposomes, we present evidence to show that 
these are also captured intact by hm-chitosan. In this experiment, we prepared liposomes 
containing the hydrophilic dye CF. We ensured that the dye was present only inside 
liposomal cores; free (unencapsulated) dye was removed via size-exclusion 
chromatography, as described in the Experimental Section. The biopolymers were left 
untagged for this experiment to avoid overlap with the green fluorescence from the CF. 
Again, hm-chitosan was deposited on the left electrode and chitosan on the right. The 
chip was then incubated with the CF-filled liposomes followed by rinsing with DI water. 
At this stage, Figure 5.6a shows a strong green signal only on the hm-chitosan side, 
 97 
implying that the liposomes preferentially attached there. The only alternate explanation 
is that the fluorescence comes from free CF dye stuck to the hm-chitosan, i.e., that the 
liposomes had already been disrupted. To rule out this possibility, we added the detergent 
Triton X-100 into the solution above the chip. This detergent is known for its ability to 
disrupt liposomes into mixed micelles, whereupon dye in the liposomal cores would be 
released into solution (this is illustrated by the schematics in Figure 5.6). After detergent 
addition and subsequent rinsing, Figure 5.6b shows that the green signal has vanished 
from the hm-chitosan film. The loss of signal after detergent treatment necessarily 
implies that the liposomes were initially intact on the hm-chitosan surface in Figure 5.6a.  
 98 
 
Figure 5.6: Evidence for intact capture of liposomes on hm-chitosan films. (a) Green-
 filtered fluorescence image showing preferential binding of liposomes (filled with the 
hydrophilic dye CF) to the hm-chitosan (H) side relative to the chitosan (C) side. The 
chip is then exposed to the detergent Triton X-100 and rinsed. The detergent disrupts the 
liposomes into micelles, as shown by the schematics, and the encapsulated dye is thus 
released and washed away. (b) Fluorescence image of the rinsed chip shows nearly 
complete loss of fluorescent signal, confirming disruption of originally intact liposomes.  
 
Further evidence for the intact capture of nanosized surfactant vesicles is provided 
by cryo-TEM. For these experiments, we deposited hm-chitosan directly on holey-carbon 
TEM grids. These grids were then incubated with surfactant vesicles, followed by a 
rinsing step with DI water to remove weakly adsorbed vesicles. The rinsed grids were 
 99 
maintained in an aqueous environment at room temperature. These grids were then 
prepared in the usual way for cryo-TEM (see Experimental Section) by plunging into 
liquid ethane. Observation of the vitrified specimens showed the presence of numerous 
spherical structures of 100-150 nm diameter (Figure 5.7). These are evidently the 
surfactant vesicles and they all seem whole ? no breaks or holes can be found on the 
vesicle shells. Thus, once again, the vesicles appear to have been captured intact.     
 
 
Figure 5.7: Cryo-TEM image of vesicles on an hm-chitosan film. The hm-chitosan was 
deposited on the TEM grid and then used to capture surfactant vesicles from solution. 
The image shows numerous spherical objects of ~ 100?150 nm diameter, which are 
presumably intact vesicles.  
 
 
Spatiotemporal Control of Vesicle Capture.  We now demonstrate the capability to 
create specific microscale patterns of vesicles on the chip. Towards this end, we used two  
6-electrode chips with a view towards engineering two specific patterns of vesicles.  On 
one chip we deposited hm-chitosan (H) and unmodified chitosan (C) in an alternating 
pattern, i.e., HCHCHC, on the 6 electrodes (Figure 5.8a). After incubating the film with a 
solution of DiI-tagged surfactant vesicles, the red signal indicative of vesicles is observed 
 100 
only on the hm-chitosan (H) electrodes (Figure 5.8b). Next, on the other chip, we 
deposited the polymers in an ?outside-inside? pattern, i.e., HCHHCH (Figure 5.8c). 
Incubation of the chip with the same tagged vesicles again results in a red signal only on 
the hm-chitosan (H) electrodes (Figure 5.8d). Thus, in both cases, pre-determined 
patterns of vesicles were successfully created. Such patterning can be easily extended to 
more complex designs and with finer resolution. Previous work with chitosan has 
indicated the resolution limits of electrodeposition to be on the order of 1 ?m.124 We view 
this work as a step towards the construction of ?vesicle arrays? that could prove to be a 
useful tool for probing the properties of biomolecules such as membrane proteins.  
 
Figure 5.8: Patterning of vesicles on 6-electrode chips. (a) NHS-tagged hm-chitosan (H) 
and chitosan (C) are electrodeposited in an alternating pattern, HCHCHC. (b) Following 
contact with DiI-tagged surfactant vesicles and rinsing, the red fluorescence indicative of 
vesicles is found only on the H electrodes. (c) H and C are electrodeposited in an 
?outside-inside? pattern, HCHHCH. (d) Once again, DiI-tagged surfactant vesicles are 
found anchored only on the H electrodes. 
 
Accessibility of Captured Vesicles. Lastly, for immobilized vesicles to be useful in 
biosensing or binding assays, the vesicles should be able to access ligands from the 
external solution. To evaluate this aspect, we studied the interaction between surface-
 101 
bound biotinylated liposomes and streptavidin in the surrounding solution. The 
experiment is depicted schematically in Figure 5.9a. Biotinylated liposomes were 
prepared by combining PC and PE-biot in a molar ratio of 9:1. These liposomes were 
then contacted with a chip bearing hm-chitosan (H) and chitosan (C) films on adjacent 
electrodes. The chip was then rinsed with DI water and exposed to a solution containing 
fluorescently tagged streptavidin, a protein that non-covalently binds to biotin with 
extremely high specificity. The results show a significantly higher fluorescent signal from 
streptavidin on the hm-chitosan electrode than the chitosan electrode (Figure 5.9b). The 
signals are quantified in the plot shown in Figure 5.9c ? the fluorescence is 
approximately 6 times higher on the hm-chitosan  side (9.3 ? 0.8 gray value) relative to 
the chitosan side (1.6 ? 0.3 gray value) . The high signal on the hm-chitosan side is 
evidently due to the binding of streptavidin to the biotinylated liposomes captured on that 
film. The binding of streptavidin to the chitosan side is not zero presumably because 
streptavidin (a negatively charged protein) can bind non-specifically to the positively 
charged chitosan. Nevertheless, this is an encouraging result because it does confirm the 
ability of immobilized vesicles to sense reporter molecules from the surrounding 
environment.  
 
 
 
 
 102 
  
Figure 5.9: Protein sensing via vesicles anchored to an hm-chitosan film. (a) Schematic 
of the experiment: biotin-decorated liposomes are first immobilized, then the chip is 
contacted with the fluorescently labeled protein, FITC-streptavidin. The binding of the 
protein is then detected by fluorescence microscopy. (b) Fluorescence image showing a 
strong signal from the hm-chitosan (H) electrode compared to the chitosan (C) electrode. 
The strong signal is indicative of protein binding to the biotins on the anchored liposomes. 
(c) Analysis of the intensities in (b). The average gray values for each electrode area are 
listed above the corresponding regions. 
 
5.4 Conclusions 
We have shown that vesicles and liposomes can be captured on electrodeposited films of 
hm-chitosan. Hm-chitosan is significantly superior in its capability for vesicle capture 
compared to unmodified chitosan, which implies that vesicle capture on hm-chitosan is 
promoted by the non-covalent binding of polymer hydrophobes to the hydrophobic 
interiors of vesicle bilayers. We have presented a range of evidence to show that the 
vesicles bound to hm-chitosan are intact structures. The evidence includes direct 
visualization of captured vesicules ? by optical microscopy in the case of microscale 
GUVs and by cryo-TEM in the case of nanoscale vesicles. Additionally, key indirect 
 103 
evidence for vesicle intactness comes from a study on dye-filled vesicles ? upon 
detergent treatment, a loss of fluorescence from the film is observed, which can only be 
attributed to the detergent-induced disruption of intact vesicles. We have also confirmed 
the ability of biotinylated vesicles bound to hm-chitosan to conjugate with the protein 
streptavidin from the surrounding solution. A particular utility of using hm-chitosan as a 
?soft? interconnect between ?hard? surfaces and vesicles is the prospect of creating 
patterns of immobilized vesicles. It is straightforward to electrodeposit hm-chitosan in a 
variety of predesigned patterns with micron-scale resolution; subsequently, vesicles can 
be readily captured on the patterned regions without the need for further chemistry (i.e., 
solely by non-covalent hydrophobic interactions). The simplicity of this approach should 
make it attractive for researchers to create new biosensors and bioassays using vesicles.  
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 104 
Chapter 6 
Gelling Blood with an Amphiphilic Biopolymer: A 
Route to Improved Hemorrhage Control 
 
6.1 Introduction 
The blood coagulation cascade is an enormously complex orchestration of small 
molecules, proenzymes, proteins and cells that ultimately results in the self-assembly 
(?polymerization?) of fibrinogen molecules into an elastic fibrin network.135 Fibrinogen 
is a small globular protein, and its assembly into fibrin chains is catalyzed by the protein 
thrombin. The fibrin network forms the hemostatic ?plug? or the physical barrier that 
prevents further loss of freely-flowing blood from the site of injury. The coagulation  
cascade is effective at arresting blood loss from small wounds, but when the injuries are 
severe, such as gunshot or knife wounds suffered by soldiers in the battlefield, alternate 
tools and strategies are required for the achievement of hemostasis.136-137 Additionally, 
there are many types of coagulopathic conditions (hemophilia, Von Willebrand disease, 
liver disease) that require medical treatment even for minor bleeding.138-140 
 
Although the need for better hemostats has existed throughout human history, it 
was not until the mid-twentieth century that major advances in the field began to develop 
beyond standard cotton gauze or torniquets. During World War II, fibrin-based materials 
(glues, foams, adhesives) were some of the first proposed materials for advanced 
hemorrhage control.141 These products generally contain extremely high concentrations 
 105 
of fibrinogen and thrombin, which work to mimic the final stages of the natural 
coagulation cascade. Early versions of these materials often demonstrated negative side 
effects such as immunogenic responses or disease transmission;142 however, improved 
processing techniques allowed for a great deal of the advancement in fibrin-based 
hemostats in the early 1990s.143-144 The approach during that period began to shift toward 
a dry powdered mixture of human fibrinogen and thrombin packed onto a solid bandage 
backing.145 By firmly pressing this dressing onto a bleeding injury, the bandage instantly 
forms a strong fibrin seal at the point of injury as bodily fluid brings the powder into 
solution in high concentration. Despite the efficacy of the fibrin sealant (FS) bandage, 
practical issues have severely limited its contributions to trauma medicine. Specifically, 
human fibrinogen and thrombin are highly expensive molecules that are scarce in supply. 
As such, they are extremely costly and largely unable to meet clinical demand.141,146 
 
Over the last decade, a myriad of other solutions have been posed to the problem 
of severe hemorrhage.147-148 Many of these employ biomolecules involved in the clotting 
cascade (e.g.  factor VIIa, Factor VIII, factor XIII, prothrombin complex concentrates). 
Unfortunately, these approaches remain costly and some have been shown to evoke 
immunogenic responses.149 Other solutions involve materials that work by means other 
than self-assembly. First in this category are powders (zeolite or mineral) that rapidly 
absorb fluid and, as a result, concentrate clotting factors at the site of injury.150 Secondly, 
there are bandages (chitosan, cellulose, gelatin) that help to either initiate clotting, absorb 
fluid, provide backpressure against flow of blood, or some combination thereof. However, 
all of these lower-cost solutions have drawbacks of either inadequate effectiveness or 
 106 
dangerous side effects (e.g. tissue/nerve damage and embolism).151-152 Furthermore, they 
all have demonstrated a great deal of user variability, particularly in uncontrolled 
environments where they are most needed.152  
 
One research group, Ellis-Benhke et al., has employed the concept of self-
 assembly, independent of the natural coagulation cascade, in order to achieve 
hemostasis.153 They used an amphiphilic synthetic peptide (RADA16) which is able to 
quickly self-assemble into nanofibers upon their introduction into biological solution.154 
While this system has only displayed efficacy in small animal models, the benefits of 
using a self-assembled material are displayed clearly in their study. Allowing the material 
itself to build a hemostatic barrier in a ?bottom-up? fashion, from nano- to macro-scale, is 
an approach likely to eliminate the user variability that plagues other proposed solutions 
to hemorrhage control.152 Of course, for practical application, such a material should be 
low cost and easily produced, which are challenging issues for these synthetic peptides.  
 
In this Chapter, we describe the use of an amphiphilic biopolymer that can 
transform liquid blood into a gel, and thereby serve as a hemostatic sealant even for 
severe wounds in large animals. The native biopolymer is chitosan, which is a material 
already of great interest in the field of wound care due to its inherent anti-microbial 
properties,155-156 its biocompatibility157 and wide availability from natural sources.158 We 
modify the backbone of chitosan with a limited number of hydrophobic tails (2.5 mol % 
of available amines), so as to create a ?comb-graft? associating polymer that remains 
soluble in water (schematic in Figure 6.1). While native chitosan has been used in several 
 107 
applications as a hemostatic material,159-162 this hydrophobic modification greatly 
enhances the capability of chitosan to control hemorrhage. Like fibrinogen 
polymerization and nanofiber assembly via the RADA16 peptide, the mechanism of 
action for this hydrophobically-modified (hm) chitosan also occurs via self-assembly; 
however, in this case, the blood cells take an active role in the formation of gel networks, 
as opposed to passively filling space in between actively polymerizing chains. Based on 
our previous work on hm-chitosan with vesicles99 (self-assembled nano-containers, which 
can be conceptualized as ?empty? cells), we hypothesize that the networks are formed by 
insertion of the hydrophobes into cell bilayers and subsequent bridging of cells by the 
biopolymer. This hypothesis is outlined in Figure 6.1. We probe the hemostatic 
mechanism of hm-chitosan via studies on blood using dynamic and steady-shear rheology, 
and by application to small (murine) and large animal (porcine) injury models.   
 
 
 
Figure 6.1. Schematic of hydrophobically-modified (hm)-chitosan as a self-
 assembling hemostat. When adding hm-chitosan to a whole blood cell suspension, the 
resulting mixture self-assembles into a 3-dimensional network. This self-assembly is 
driven by insertion of hydrophobes into blood cell membranes and the subsequent 
physical crosslinking of adjacent cells. 
 108 
6.2 Experimental Section 
Materials. Chitosan of medium molecular weight (190-310K) and Brookfield viscosity 
of 286 cps was obtained from Sigma-Aldrich. The reported degree of deacetylation was 
about 80%. Chitosan was dissolved in 0.15 M D-Lactic Acid (LA) so as to mimic 
physiological ionic strength. D-Lactic Acid and sodium cyanoborohydride were 
purchased from Sigma-Aldrich. 4-octadecylbenzaldehyde and ?-cyclodextrin were 
purchased from TCI International. Hank?s Balanced Salt Solution (HBSS) was purchased 
from Lonza. BD Vacutainers? were purchased from Becton Dickinson.  
 
Synthesis of Chitosan Derivatives. The hm-chitosan was synthesized by attaching 
benzene-n-octadecyl tails to the chitosan backbone via reaction with 4-
 octadecylbenzaldehyde. The same procedure, but instead using a dodecylaldehyde as the 
reactant, has been reported in our earlier paper99 and it also follows those described in the 
literature.17,98,131 The degree of hydrophobic substitution follows the reaction 
stoichiometry and in this study it was fixed at ca. 2.5 mol% of the available amine groups. 
The degree of hydrophobic substitution follows the reaction stoichiometry and in this 
study it was fixed at ca. 2.5 mol% of the available amine groups.  
 
Manufacture of hm-chitosan Sponges. 0.5 wt% hm-chitosan in 0.15 M Lactic Acid was 
poured into trays of dimensions 10 cm (length) ? 10 cm (width) ? 4 cm (height). Trays 
were filled only up to 2 cm in height with polymer solution. Samples were then frozen to 
-40?C in a pilot-scale freezed drier, and then placed under vacuum of 50 ?bar for a period 
of 5 days in order to remove all water. After drying, a Tegaderm? medical tape backing 
 109 
was placed on the back side of the sponges for unhindered compression of the sponge 
during surgery. 
 
Obtaining Blood. 5 human subjects volunteered to have 20 ml of blood drawn by 
registered nurse at the University of Maryland School Medicine. Subjects were healthy 
adults ranging age from 20 to 40 years of age (4 males, 1 female). 10 ml intervals of  
blood were drawn into BD Vacutainers? containing 143 USP units of Sodium Heparin. 
The protocol was approved by the Institutional Research Board (IRB) at the University of 
Maryland.  
 
Rheology of Chitosan-Blood Mixtures: Steady and dynamic rheological experiments 
were performed on a Rheometrics AR2000 stress-controlled rheometer. A cone-and-plate 
geometry of 40 mm diameter and 4? cone angle was used and samples were run at 
physiological temperature 37?C. Dynamic frequency spectra were obtained in the linear 
viscoelastic regime of the samples, as determined by dynamic strain sweep experiments. 
 
Murine Injury Models. Surgical procedures were approved by the Institutional Animal 
Care and Use Committee of the University of Maryland. Fasted male Long-Evans rats (n 
= 15, 250?275 g; Harlan, Indianapolis, IN) were anesthetized (60 mg/kg ketamine and 7.5
 mg/kg xylazine given IP) and allowed to breathe air spontaneously. Animals were 
maintained under pathogen-free conditions in 12 h diurnal cycles, with water and food ad 
libitum. Animal rooms were kept at 21?3 ?C with several changes of air per hour. All 
husbandry and animal procedures were in accordance of humane animal handling 
 110 
practices under the guidance of the Unit for laboratory Animal Medicine at the University 
of Maryland School of Medicine. At the end of each procedure, all animals were 
humanely sacrificed by ketamine administration.  
 
Using a scalpel, the femoral vein was transected and allowed to bleed for 30 
seconds, after a unilateral groin incision was made over the femoral canal. Exposure and 
isolation of at least 1 cm of the femoral vein was performed. 1 ml of test material was 
then be dispensed onto 5 randomly selected animals via syringe (with 22 G needle) after 
wiping away excess blood from the site of injury via cotton gauze. Bleeding time was 
measured via stopwatch with the start time corresponding to the application of sample 
and end time corresponding to visual observation of halted bloodflow. Test materials 
studied were (1) Saline Buffer, (2) 0.5 wt% Chitosan and (3) 0.5 wt% hm-chitosan. 
 
Pig Injury Models. 3 Yorkshire crossbred swine, age 2.5 months and weighing 39.6 kg-
 42.8 kg, were used. A veterinarian screened all animals. Animals were allowed free 
access to water and to commercial food, which was withheld the night before the study. 
All animals were maintained in an Association for Assessment and Accreditation of 
Laboratory Animal Care International?accredited facility, and all experimental 
manipulations were performed in accordance with the National Research Council?s Guide 
for the Care and Use of Laboratory Animals. The protocol was approved by the 
Institutional Animal Care and Use Committee. The swine were anesthetized with 6 
mg/kg of Telazol (Fort Dodge Animal Health, Fort Dodge, IA, USA) and 0.01 mg/kg of 
glycopyrrolate intramuscularly. They were intubated and placed on mechanical 
ventilation at a tidal volume 12 mL/kg, a rate of 10 respirations per minute, and 100% 
 111 
oxygen. Anesthesia was maintained using isoflurane and ventilatory parameters were 
maintained to attain an end-tidal pCO2 of 40 mm Hg. 
 
The animals underwent midline laparotomy, and a unilateral groin incision was 
made over the femoral canal. Exposure and isolation of at least 5 cm of the femoral artery 
was performed. Using a 4.4 mm aortic punch, the femoral artery was punctured and free 
bleeding was allowed for 45 seconds. Blood loss was collected by suction for this period 
and was designated pretreatment total blood loss. Dry hm-chitosan sponges (bandages 
were cut to optimal sizes for wound approach) were applied via 2 min direct compression 
onto the injury site. Animals were observed under anesthesia for 180 minutes, at which 
point they were then be euthanized with Pentobarbital IV 100-200 mg/kg.   
 
 112 
6.3. Results and Discussion 
 
Figure 6.2. Comparison of the native and hydrophobically-modified polymers with 
regard to their physical influence on blood. In (a), photographs show hm-chitosan at 
0.25 wt% mixed with heparinized human whole blood (50 wt %) is able to hold its 
weight upon test tube inversion, whereas unmodified chitosan at 0.25 wt% mixed with 
heparinized human whole blood (50 wt %) remains as a liquid. A cartoon of the proposed 
?bridging? effect of hm-chitosan on blood cells is shown above the left photograph; a 
contrasting schematic of unmodified chitosan mixed with blood is above the right 
photograph. In (b), steady shear rheology displays a dramatically increased (~O(105)) 
zero-shear viscosity of the hm-chitosan-blood mixture (red circles), relative to both hm-
 chitosan (0.25 wt%) with no blood (cyan hexagons) and chitosan-blood (yellow triangles) 
controls. In (c), dynamic rheology of the hm-chitosan blood mixture (G?, closed red 
circels, G? closed green triangles) is that of gel-like elastic material which relaxes slightly 
over time, whereas rheology of the unmodified chitosan/blood mixture (G?, open green 
triangles, G?, open red circles) is that of a viscous sol 
 
 113 
Results on Gelling Blood. Figure 6.2 compares hm-chitosan and unmodified chitosan 
with regard to their affect on a heparinized (14.3 units/mL) suspension of human whole 
blood. First, we note that visual observation of these samples show the blood mixed with 
hm-chitosan holds it own weight upon vial inversion (Figure 6.2a). A similar visual  
result has been observed in a mixture of citrate-stabilized blood and a polyethyleneoxide 
(PEO) polymer modified with cholesterol groups at either end.163 In contrast, unmodified 
chitosan at the same concentration mixed with heparinized human whole blood (50 wt %) 
remains as a freely flowing liquid. A cartoon of the proposed ?bridging? effect of hm-
 chitosan on blood cells is shown above the left photograph; a contrasting schematic of 
unmodified chitosan mixed with blood is above the right photograph. In Figure 6.2b, 
steady shear rheology displays a dramatic (~ million-fold) increase in the zero-shear 
viscosity of the hm-chitosan-blood mixture, relative to both hm-chitosan (0.25 wt%) with 
no blood (cyan hexagons) and chitosan (0.25 wt%)-blood (yellow triangles) controls.  
 
In Figure 6.2c, dynamic rheology gives us information about the microstructure of 
these samples. Note that the key parameters given by dynamic rheology are the elastic 
modulus, G? (i.e. the elastic or solid like character of the material) and the viscous 
modulus, G?, (i.e. the viscous or liquid-like character of the material). Both moduli are 
measured as a function of oscillatory shear frequency, and as a result, give us information 
about the viscoelastic character over a wide range of timescales. Dynamic rheology of the 
hm-chitosan blood mixture is that of a gel-like elastic material with G? higher than G? 
over the range of practical frequencies (100 rad/s to 0.1 rad/s) and with both moduli 
showing a weak frequency dependence. No crossover of G? and G? is observed, 
 114 
indicating that the sample does not relax over long time scales (this is consistent with the 
sample-spanning network structure shown in Figure 6.2a).164-165 In contrast, rheology of 
the unmodified chitosan/blood mixture (G?, open green triangles, G?, open red circles) is 
that of a viscous sol. That is, in this case, G? exceeds G? over the frequency range and 
both moduli are strong functions of frequency.  
 
 
 
Figure 6.3. Time-to-gelation of hm-chitosan/blood mixtures. In (a), a time-sweep of 
an hm-chitosan (0.25%)/blood (50%) mixture displays the characteristics of an elastic 
material (G? [red triangles] > G? [green triangles]) over the studied interval of 11 min. 
G? > G? immediately at t = 0 min for this sample. In (b), unmodified chitosan 
(0.25%)/blood (50%) mixture shows that the sample remains as a viscous sol (G? [green 
triangles] > G? [red triangles]) over the same interval. Lastly, in (c), a sample comparison 
in a rat femoral vein injury model demonstrates that hm-chitosan (HM-CS) solution 
(0.5%) significantly decreases bleeding time relative to saline buffer (SAL) and native 
chitosan (CS) solution  (0.5%) controls. 
 
 115 
 
In Figures 6.3a and 6.3b, we probe the issue of time to gelation. This is again a 
rheological comparison between hm-chitosan/blood and chitosan/blood mixtures using a 
time sweep at a constant frequency of 10 rad/s. The experiments were done by mixing a 
given polymer and blood and quickly loading into the rheometer. For the hm-
 chitosan/blood mixture, G? exceeds G? immediately at t = 0, and thereafter there is a 
small increase in G? over the studied time interval of 11 min. In contrast, the 
unmodified/blood mixture displays much lower values of G? and G?, with G? exceeding 
G? throughout the studied time interval of 11 min. This suggests that hm-chitosan gels 
blood rapidly upon application. It also highlights the fact that the hydrophobes grafted to 
the chitosan backbone significantly enhance the hemostatic ability of the polymer. 
 
Results from Animal Injury Models. Figure 6.3c displays a bleeding time study on a 
minor injury model in a small animal. Femoral vein injuries were created in Long-Evans 
adult rats (n = 5 per sample) via scalpel. Bleeding times were measured for all injuries. 
All materials were applied to the injury via a 22 G needle syringe. One mL of a saline 
control achieved hemostasis in 50 ? 4 s; an equal volume of an unmodified chitosan 
control (0.5 wt%) showed a similar time to hemostasis of 47 ? 3 sec. In contrast, 1 mL of 
an hm-chitosan solution, also at 0.5 wt%, achieved hemostasis in a period of 4.5 ? 0.6 sec. 
The hm-chitosan was also able to control bleeding from a porcine femoral vein injury in a 
comparable period of time (5.6 ? 0.7 s). Hence, this amphiphilic chitosan displays 
promise over unmodified chitosan as a hemostat for minor wounds in animals.  
 
 116 
In order to treat high-pulsatile injuries in large animals, we produced freeze-dried 
bandages of this hm-chitosan. 150 mL of a 0.8 wt% hm-chitosan solution was prepared 
and then cast into a plastic container of dimensions 4? ? 4? ? 0.5?. The solution was 
frozen at ?40?C and vacuumed at 50 ? 10-6 mbar for several days.  Similar bandages of 
unmodified chitosan have been used commercially, particularly for US troops in the 
conflicts in Iraq and Afghanistan.146,166 While these unmodified chitosan bandages 
generally provide adequate initial first aid, they fail to maintain hemostasis for a period of 
longer than 1.6 h, and they allow secondary bleedings to occur.167 This is likely due to 
loss of tissue adhesion during the studied timeframe as the bandages become saturated 
with liquid blood, thus displacing key electrostatic interactions between chitosan and soft 
tissue.  
 
We have performed an evaluation of the effectiveness of the hm-chitosan 
bandages on a pig femoral artery injury model (n = 3). Figure 6.4 displays photographs 
from a representative application of the bandage to a femoral artery puncture (4.4 mm 
punch), which is fatal within 15 minutes left untreated. Figure 6.4a displays the blood 
pooling with the open cavity after puncture, Figure 6.4b displays the application of the 
bandage, Figure 6.4c displays the bandage successfully halting bleeding after 2 min of 
compression, and Figure 6.4d shows the arterial puncture successfully clotted after 
removal of the bandage (3 h after application). All pigs survived for the duration of the 
experiment, after which they were euthanized according to protocol. In sum, the bandage 
was highly effective at treating a severe bleeding injury in pigs, no secondary bleedings 
occurred during the studied timeframe, and no toxicity was observed.  
 117 
 
 
 
Figure 6.4. Application of an hm-chitosan sponge to a porcine femoral artery 
puncture. In (a) the injury was created in the femoral artery, resulting in high-pulsatile 
flow. An hm-chitosan is then applied to the injury in (b) by direct compression. After 2 
min of compression, the bandage is observed in (c) to be attached to the wound site and 
providing hemostasis. The bandage is removed after 180 min (d) with no rebleeding 
during the studied interval.  
 
 
We believe that a dual effect is occurring during the use of the hm-chitosan 
bandage on severe injuries. First, the hm-chitosan that comes in direct contact with the 
blood dissolves slightly, and in turn, helps gel the blood in the manner highlighted in 
Figure 6.2a. Because the blood in the vicinity of the bandage gets gelled, it does not soak 
up the bandage readily, and the gelled blood presumably acts as a barrier to blood loss 
through the puncture. (This is similar to the postulated action of solid fibrin bandages, as 
discussed in the Introduction.) Secondly, the parts of the bandage that come in direct 
contact with the tissue are able to adhere much more strongly to the tissue than an 
unmodified chitosan bandage. Indeed, in vitro tissue adhesion studies have shown that 
increasing the level of hydrophobic modification along the chitosan backbone enhances 
the peel strength of the chitosan in a linear fashion (data not shown). The improved 
 118 
adhesion decreases the likelihood of bandage delamination and thereby the possibility of 
secondary bleeding. We believe the improved adhesion of hm-chitosan to tissue may also 
be a consequence of hydrophobic interactions between polymer hydrophobes and the 
underlying tissue cells.  Although these hydrophobic interactions are non-covalent and 
hence weak individually, they sum up to a significantly large interaction over the entire 
surface area of the tissue. As such, the bandage acts like a nano-scale Velcro material in 
its interaction with soft tissue, i.e. it adheres strongly due to many nano-scale interactions, 
but yet it can be conveniently removed with no damage to underlying tissue or nerves 
since each individual interaction is weak.  
 
Probing the Mechanism for Blood Gelling. In order to probe the mechanism of gel 
formation, we separated blood plasma from the hemotacrit via centrifugation. The cells 
were resuspended in an equal volume of Hank?s Balanced Salt Solution and then mixed 
with hm-chitosan. Figure 6.5 shows the dynamic rheology of 0.25% hm-chitosan with 
50% diluted blood cell suspension. The rheology mirrors that of the whole blood mixed 
with the same concentration hm-chitosan quite closely, with G? > G? over the entire 
range of frequencies. Also, it is observed visually that the mixture, like whole blood, 
holds its own weight upon vial inversion. In contrast, the mixture of 0.25% hm-chitosan 
with 50% diluted plasma shows G? > G? over the range of frequencies, and visually 
appears to flow freely. This set of results shows that the cells play a key role in the 
gelation process.  
 119 
 
Figure 6.5. Dynamic rheology of hm-polymer mixed with separated blood 
hematocrit and plasma. The hm-chitosan concentration is maintained at 0.25%. 
Hematocrit was resuspended in isotonic buffer after centrifugation; plasma was removed 
and stored for use. Visual observation of an hm-chitosan-hemotacrit mixture shows that 
the sample can hold its weight upon vial inversion, whereas an hm-chitosan-plasma 
mixture flows freely as a viscous liquid. Dynamic rheology confirms the visual evidence, 
as the hematocrit mixed with polymer (G? [closed red circles] and G?[closed green 
circles]) is that of a gel-like elastic material which relaxes slightly over long times. In 
contrast, the rheology of the plasma mixed with polymer (G? [open red triangles] and 
G?[open green triangles]) is close to that of a sol-gel transition.  
 
Reversal of Blood Gelling. While new hemostatic technologies have advanced rapidly 
over the past 15 years, there always exists a concern of undesired embolism formation 
from a new hemostat. In our probing of the mechanism of blood gelation via hm-chitosan, 
we have incidentally discovered a means of mitigating concerns of blood vessel 
embolization. Cyclodextrins (CDs) are a class of supramolecules known for their ability 
to modulate hydrophobic interactions between associating polymers.33,38 CDs are 
 120 
biologically safe molecules often used frequently in foods, pharmaceutical excipients, etc. 
Of the three major cyclodextrins (CDs), (1) ?-CD, (2) ?-CD, and (3) ?-CD, it is ?-CD that 
shows a strong affinity for single-tailed hydrophobes. In a rheological study previously 
reported from our group, all three cyclodextrins were added to aqueous solutions of 
telechelic polyethylene oxide with single tailed n-alkyl grafts on either end.168 The ?-CD 
decreased the viscosity of the polymer solution much more rapidly than the other two CD 
species. ?-CD has also been used in several other  studies as a viscosity reducer for 
associating biopolymers. This phenomenon is thought to occur due to the molecular 
architecture of the ?-CD. ?-CD is a barrel shaped supramolecule; the exterior of the 
barrel is very hydrophilic, allowing solubility in water, whereas the center of the barrel is 
hydrophobic. The diameter of the barrel is 5.4 ?, which in contrast to that of ?-CD and 
?-CD, is optimal for associating with single-tailed alkyl chains. In this way, the barrels 
are able to ?cap? the hydrophobes along the backbone of the polymer, and hence screen 
their hydrophobic interactions from one another, in effect reducing the apparent viscosity 
of a bulk solution of hydrophobically-modified polymer. 
 
Because of ?-CD?s ability to eliminate hydrophobic interactions among 
neighboring polymer chains, we hypothesized that given enough concentration of ?-CD 
in solution, this supramolecule would be able to displace existing interactions between 
the hydrophobes and the bilayers of blood cell membranes. A schematic of this ?-CD 
?screening? hypothesis is shown above in Figure 6.6a. The key concept here is that the 
addition of ?-CD in aqueous media to an hm-chitosan/blood gel, triggers a ?self-
 disassembly? of the elastic 3-dimensional network into a freely flowing fluid. In Figure 
 121 
6.6b, dynamic rheology confirms this hypothesis as a hm-chitosan (0.25%)/blood (50%) 
displays gel-like characteristics with G? exceeding G? for the studied range of 
frequencies, whereas the same mixture hm-chitosan (0.25%)/blood (50%) with 3 wt% 
?-CD included displays the rheology of a freely-flowing liquid, with G? exceeding G? 
over the range of frequencies and significantly lower moduli values.  
 
 
 
Figure 6.6. Reversal of blood-gelation via ?-cyclodextrin (CD). In (a), a cartoon of ?-
 CD displays its supramolecular barrel-shaped structure, with hydrophobic core and 
hydrophilic exterior. A schematic hypothesis shows that by adding these molecules 
together with hm-chitosan-blood mixtures, the samples become liquefied due to 
?capping? of hydrophobes by the ?-CD molecules. In (b), dynamic rheology confirms 
this hypothesis as a hm-chitosan (0.25%)/blood (50%) displays gel-like characteristics 
(G? [closed red circles] and G?[closed green triangles]), whereas a hm-chitosan 
(0.25%)/blood (50%)/?-CD (3%) mixture displays the rheology of a freely-flowing liquid 
(G? [open red circles] and G?[open green triangles]).  
 
The results of this experiment have two important implications. The first is a 
practical one. That is, this gelation process, or ?artificial clotting,? is reversible by the 
 122 
addition of a low-cost, biocompatible molecule, ?-CD. This is a key mitigation against 
embolization potentially caused by this system. Although neither dangerous clotting nor 
toxicity were observed during our animal studies, it is useful to have the capability to 
reverse the action of a hemostatic agent. The second implication is a further insight into 
the mechanism of gelation. Because the ?-CD has been show to negate the interaction of 
the hydrophobes within associating biopolymer solutions in water, we can conclude that 
the hydrophobes must play an important role in the blood gelation process. 
 
6.4. Conclusions 
We have shown that the amphiphilic biopolymer, hm-chitosan, can act as an 
effective hemostatic agent. It has the ability to transform whole liquid blood into a gel, 
and it quickly stops bleeding from both minor and severe injuries in small and large 
animals. The parent polymer, chitosan, is itself commercially used as a material for 
hemostatic bandages, presumably because of its cationic nature and anti-microbial 
properties. However, chitosan does not change the physical character of liquid blood, i.e., 
a mixture of chitosan and blood remains a sol. In contrast, the addition of hm-chitosan 
converts liquid blood into an elastic gel with a sufficient yield stress so as to hold its 
weight in an inverted tube. The gelling mechanism evidently arises due to the 
hydrophobes on the hm-chitosan chain; we postulate that the hydrophobes anchor in 
blood cell membranes and thereby bridge the cells into a 3-dimensional network. The 
cells are thus active components (nodes or junction points) in the network rather than 
being physically trapped in a polymer mesh. The gelling ability of hm-chitosan is similar 
to that of fibrin-based sealants, but at a much lower cost and wider availability. A further 
 123 
unique aspect of hm-chitosan is that its gelling effect on blood can be reversed by 
addition of supramolecule ?-CD. The reversal occurs because ?-CD molecules sequester 
the hydrophobes and prevent them from attaching to blood cells. In sum, we suggest that 
amphiphilic biopolymers (hydrophobes attached to a hydrophilic backbone) represent a 
revolutionary paradigm in hemostatic technologies. 
 
 
 124 
Chapter 7 
Conclusions and Recommendations 
 
7.1. Conclusions 
The goal of building nano-machines for improving human health from the nano-
 scale upward is a noble and ambitious one. Progress in the area of nanomedicine will 
hinge upon a fundamental understanding and embracing of the soft and ?sticky? 
environment which exists at the biological nano-scale. Taking inspiration from the cell as 
the ultimate ?soft machine,? one which is able to work beautifully at the nano-scale, we 
have blueprinted 4 self-assembled soft-material systems which display advanced 
functionalities with applications in the areas of controlled drug delivery, tissue 
engineering, and wound care.  
 
 We have shown that nanoscale vesicles of NaOA can be entrapped within gelatin 
hydrogels. The resulting vesicle-gel hybrids exhibit the pH-responsive properties of the 
NaOA vesicles. Specifically, when exposed to a pH 10 buffer, the vesicles within the gel 
become transformed into micelles. Vesicle disruption can be done in a controlled manner 
at specific locations within a gel. Gels can thus be ?patterned? to have vesicle-rich and 
micelle-rich domains in predetermined arrangements. The utility of entrapping pH-
 responsive structures within the gel is in the area of controlled release of hydrophilic 
solutes. We show that the release of calcein dye out of a vesicle-gel into the external 
 125 
solution is accelerated when the solution pH is raised to 10. This increase is attributed to 
a pH-induced vesicle to micelle transition within the gel, which reduces the transport 
resistance to dye diffusion.    
 
 We have shown that ?mothership? capsules with hierarchical structural features 
can be formed by adding droplets of a feeder solution consisting of liposomes and 
chitosan, a cationic biopolymer, into a solution of anionic biopolymer (gellan gum) or 
surfactant (SDBS). The resulting structures are spherical capsules; assembly is achieved 
via electrostatic interactions between oppositely charged polymers/surfactants at the 
interface of the droplet. Capsule size is simply dictated by drop size. Chitosan-gellan 
motherships are able to sustain model drug release due via intact liposomes carrying the 
drug within the capsule. Chitosan-gellan motherships can be degraded by the addition of 
the enzyme chitosanase; complete degradation occurs over a 7 period, with intact vesicles 
present in the remaining solution post-degradation. Finally, hierarchical capsule-within-
 capsule structures were produced with either chitosan-SDBS or chitosan-gellan systems.  
 
 We have shown that vesicles and liposomes can be captured on electrodeposited 
films of hm-chitosan. Hm-chitosan is significantly superior in its capability for vesicle 
capture compared to unmodified chitosan, which implies that vesicle capture on 
hm-chitosan is promoted by the non-covalent binding of polymer hydrophobes to the 
hydrophobic interiors of vesicle bilayers. We have presented a range of evidence to show 
that the vesicles bound to hm-chitosan are intact structures. The evidence includes direct 
visualization of captured vesicules ? by optical microscopy in the case of microscale 
 126 
GUVs and by cryo-TEM in the case of nanoscale vesicles. Additionally, key indirect 
evidence for vesicle intactness comes from a study on dye-filled vesicles ? upon 
detergent treatment, a loss of fluorescence from the film is observed, which can only be 
attributed to the detergent-induced disruption of intact vesicles.  
 
Finally, we have shown that hm-chitosan, an amphiphilic biopolymer, can act as 
an effective hemostatic agent. It has the ability to transform whole liquid blood into a gel, 
and it quickly stops bleeding from both minor and severe injuries in small and large 
animals. The parent polymer, chitosan, does not change the physical character of liquid 
blood, i.e., a mixture of chitosan and blood remains a sol. The gelling mechanism 
evidently arises due to the hydrophobes on the hm-chitosan chain; we postulate that the 
hydrophobes anchor in blood cell membranes and thereby bridge the cells into a 3-
 dimensional network. A further unique aspect of hm-chitosan is that its gelling effect on 
blood can be reversed by addition of supramolecule ?-CD. The reversal occurs because 
?-CD molecules sequester the hydrophobes and prevent them from attaching to blood 
cells.  
 
7.2. Recommendations for Future Work 
Spatio-temporal control of the transition of fatty acid vesicles to micelles with the 
gels presentated in Chapter 3 would have some interesting implications. For example, we 
could make potential build different gelatin gel ?sections? adjacent to one another that 
contain different types of model drugs. It would be nice to have a way to selective choose 
when to release a certain kind of drug into a surrounding buffer solution. This would be 
 127 
feasible with a spatiotemporal pH-trigger. There is a commercially available photobase 
generator called MGC which produced OH- ion upon UV irradition.169 By incorporating 
this molecule into the bilayer of the vesicles prior to embedding them within the gels, we 
could subsequently use UV lasers to point at different spots within the gels where we 
want a vesicle-to-micelle transition to occur. A schematic of MGC generation of 
hydroxide ion is show in figure 5.1 below: 
 
Figure 7.1. Photolysis of MGC, a photobase generator (PBG). Upon UV irradiation, the 
molecule dissociates and releases OH?.  
 
The ?mothership? systems described in Chapter 4 have some interesting 
implications with regard to developmental biology or origin of life studies. For example, 
a water-soluble glycol chitosan could be dissolved in cell culture media along with a 
suspension of cells, for example, Human Umbilical Vein Endothelial Cells (HUVECs). 
Another set of capsules could contain a suspension of proteins, Vascular Endothelial 
Growth Factor. This would allow us to do very simple, but intriguing experiments where 
we could take one capsule containing cells and another capsule containing growth factor, 
then place the two within the same capsules via the one-step drop capsule-within-capsule 
method described in Chapter 4. It is possible that the cells may be able to escape their 
capsule and form a primitive blood vessel in the vicinity of the VEGF capsule. This 
 128 
hypothelial scenario expresses the type of isolated signaling that can take place during 
embryo formation or tumor growth. More generally, a range of different packaging and 
release experiments could be done with these systems, and many could yield interesting 
results. 
 
We remain on the subject of growth factors with regard to future work on the 
vesicle-biopolymer films described in Chapter 5. Certain growth factors, such as platelet-
 derived growth factor (PDGF) and epidermal growth factor (EGF), have been show to 
accelerate the healing of chronic wounds, however their bolus administration can be 
problematic and costly.170 The vesicle hm-chitosan films provide a simple way to 
therapeutically functionalize the films with both of these growth factors, and presumably 
would release the factors in a sustained fashion in aqueous media. (1) Proteins could be 
packaged inside the vesicles, (2) the vesicles could then be attached to the films, (3) and 
the films could then be removed from the electrodes and (4) finally placed in buffer so as 
to test the release of protein. Additionally, it would be worth making these films with the 
active component inside the vesicle and try out the films on chronic wounds in diabetic 
mice, relative to unmodified chitosan controls and bolus PDGF. Any progress in that 
regard could be very exciting. 
 
With regard to the use of amphiphilic polymers for gelling blood, many more 
experiments can and should be tried. It would be instructive to try out a range of 
amphiphilic polymers, e.g. hm-alginate, hm-gelatin, hm-hyaluronic acid, etc., on the 
blood samples to determine if there are a few samples that work particularly well with 
 129 
regard to gel strength and gelation time. These ?winners? could then move on to small 
and large animal studies in head-to-head tests with fibrin-based products which dominant 
much of the clinical hemostat space. Of course, more large animal studies should be 
completed for the bandages that were presented in pig models in Chapter 6 and some key 
controls should be tested in those models. On a grander scheme, it would interesting to 
try to correlate the rheology of a given hemostat mixed with heparinized blood in vitro 
with the performance of the hemostat in important in vivo bleeding models. Because 
rheology experiments are much less expensive than animal studies, the existence of any 
positive correlation between in vitro and in vivo could be very valuable. 
 
Lastly, it would be interesting to develop an hm-chitosan which is able to dissolve 
in cell culture media at pH 7.4, so that it could potentially gel healthy soft tissue cells. In 
this way, the hm-chitosan could function as a useful delivery vehicle for cells used as 
therapeutics (e.g. stem cell therapy for damaged heart tissue). ?Cellular therapy? refers to 
the use of cells themselves as therapeutic agents, as opposed to small molecules or 
proteins as the sole active agent. A major drawback in the field of cellular therapy is lack 
of effective delivery mechanisms for the cells, and an associating biopolymer could 
potentially fill this void.171 
 
 
 
 
 
 130 
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