EFFECT OF NANOSCALE AND HIERARCHICAL TOPOGRAPHIES ON THE ANTIFOULING EFFICACY OF SILICONE SURFACES
By
CLAYTON WALKER ARGENBRIGHT
A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA
2018
© 2018 Clayton Walker Argenbright
To my family
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ACKNOWLEDGMENTS
I would like to thank my advisor Dr. Anthony Brennan who has constantly
supported me throughout graduate school. I also must thank my committee members
for access to their expertise and laboratories, without which this work would not have
been possible: Dr. Scott Perry, Dr. Christopher Batich, Dr. Thomas Angelini, and Dr.
Antonio Webb. Dr. Perry was also my advisor and employer during my undergraduate
studies, giving me my first taste of research and a way to support myself financially
while focusing on my degree. I would not be here today without your help. This work
would also not be possible without the help of our collaborators Dr. Anthony Clare and
Dr. John Finlay, who performed all the biofouling assays in this study.
I cannot possibly overstate the value of the help received from former and current
group members of the Brennan Research Group: Dr. Joseph Decker, Dr. Canan
Kizilkaya, Dr. Laura Villada, Dr. Cary Kuliasha, Mr. Francisco Castro-Cara, Dr. Ha
Nguyen, Mr. Vignesh Nandakumar, and Mr. Yi Wei. Other UF graduate students and
research groups have also been extremely helpful, most notably Mr. Sin-Yen Leo and
Dr. Peng Jiang of The Jiang Group. I would also like to thank Dr. Jon Dobson and his
research group for their scientific expertise and moral support on campus, and their
throwing arms on the field.
None of this work could have been accomplished without the excellent staff of
the many UF facilities used, like MAIC and the NRF. Dr. Brent Gila, Mr. Eric Lambers,
Mr. Bill Lewis, Mr. Al Ogden, and Dr. David Hays; thank you for your time, patience, and
everything you have taught me.
Most importantly, I must thank my whole family as well who has supported and
encouraged me for my entire life.
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TABLE OF CONTENTS page
ACKNOWLEDGMENTS .................................................................................................. 4
LIST OF TABLES ............................................................................................................ 8
LIST OF FIGURES .......................................................................................................... 9
LIST OF ABBREVIATIONS ........................................................................................... 11
ABSTRACT ................................................................................................................... 13
CHAPTER
1 INTRODUCTION .................................................................................................... 15
Scope of Research ................................................................................................. 15
Specific Aims .......................................................................................................... 15 Biofouling ................................................................................................................ 16
Antifouling Strategies .............................................................................................. 19 Non-toxic Antifouling Strategies .............................................................................. 21 Antifouling Surface Topographies ........................................................................... 22
Wetting Behavior .................................................................................................... 22 Conclusion .............................................................................................................. 25
2 ADDITION OF NANOTOPOGRAPHY TO ANTIFOULING SILICONE MICROTOPOGRAPHY ........................................................................................... 26
Background ............................................................................................................. 26 Microtopographies ............................................................................................ 26
Hierarchical Topographies ................................................................................ 28 Objectives ............................................................................................................... 29 Materials ................................................................................................................. 29 Methods .................................................................................................................. 30
Microscale Patterning ....................................................................................... 30 Langmuir-Blodgett Coating ............................................................................... 31
Reactive Ion Etching ........................................................................................ 31 HMDS Treatment ............................................................................................. 32 Polymer Replication ......................................................................................... 32 Sample Mounting ............................................................................................. 33 Characterization ............................................................................................... 33
Ulva linza Settlement Assay ............................................................................. 34 Results and Discussion........................................................................................... 34
Wafer Mold Fabrication .................................................................................... 34 Polymer Replication ......................................................................................... 35 Ulva linza Settlement Assay ............................................................................. 36
6
Conclusion .............................................................................................................. 37
3 EFFECT OF POLYDIMETHYLSILOXANE NANOTOPOGRAPHIES ON THE SETTLEMENT OF THE SPORES OF Ulva linza ALGAE ....................................... 43
Background ............................................................................................................. 43 Nontoxic Antifouling Strategies ......................................................................... 43 Nanotopographies ............................................................................................ 44
Objectives ............................................................................................................... 46
Materials ................................................................................................................. 46 Methods .................................................................................................................. 47
Langmuir-Blodgett Coating ............................................................................... 47 Reactive Ion Etching ........................................................................................ 47
Polymer Replication ......................................................................................... 47 Characterization ............................................................................................... 48
Ulva linza Settlement Assays ........................................................................... 48 Results and Discussion........................................................................................... 48
Nanotopographies ............................................................................................ 48
Polymer Replication ......................................................................................... 50 Assay 1............................................................................................................. 51
Assay 2............................................................................................................. 51 Conclusion .............................................................................................................. 52
4 SYLGARD 184 NANOTOPOGRAPHIES FOR ANTIFOULING SILICONE SURFACES ............................................................................................................ 57
Background ............................................................................................................. 57 Objectives ............................................................................................................... 57 Materials ................................................................................................................. 58
Methods .................................................................................................................. 58 Polymer Replication ......................................................................................... 58
Bioassay Sample Preparation .......................................................................... 59 Ulva linza Settlement Assay ............................................................................. 60
Results and Discussion........................................................................................... 60 SR 415 Molds ................................................................................................... 60
Sylgard 184 Topographies ............................................................................... 61 Ulva linza Settlement Assay ............................................................................. 61
Conclusion .............................................................................................................. 62
5 MECHANICAL PROPERTIES OF PDMSe SURFACES......................................... 66
Background ............................................................................................................. 66 Objective ................................................................................................................. 67 Materials ................................................................................................................. 67
Methods .................................................................................................................. 68 Results and Discussion........................................................................................... 68 Conclusion .............................................................................................................. 69
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6 MICROSCALE WETTING BEHAVIOR ................................................................... 72
Background ............................................................................................................. 72
Materials ................................................................................................................. 73 Methods .................................................................................................................. 73 Results .................................................................................................................... 74 Conclusion .............................................................................................................. 75
7 NANOTEMPLATING WITH BLOCK COPOLYMERS ............................................. 81
Background ............................................................................................................. 81 Materials ................................................................................................................. 82 Methods .................................................................................................................. 83
Solvent Casting Thick Films ............................................................................. 83 Spin Coating Thin Films ................................................................................... 83 Etching ............................................................................................................. 84
Characterization ............................................................................................... 84 Results .................................................................................................................... 84
Solvent Casting ................................................................................................ 84
Spin Coating ..................................................................................................... 85 Etching ............................................................................................................. 85
Conclusion .............................................................................................................. 86
8 SUMMARY AND FUTURE WORK ......................................................................... 88
LIST OF REFERENCES ............................................................................................... 90
BIOGRAPHICAL SKETCH ............................................................................................ 96
8
LIST OF TABLES
Table page 3-1 RIE etching parameters for nanopatterned wafers. ............................................ 53
9
LIST OF FIGURES
Figure page 2-1 Two dimensional and three dimensional representations of silica particle
monolayer on wafer surface imaged with AFM in ScanAsyst mode. .................. 38
2-2 SEM micrograph of the fracture surface of a silicon wafer with nanotopography etched into the surface. ........................................................... 38
2-3 SEM micrographs of nanotopography etched into silicon wafer surface ............ 39
2-4 Average contact angle data from water droplets on wafer and PDMSe surfaces throughout replication process ............................................................. 40
2-5 SEM micrograph of part of a unit cell of Sharklet AF microtopography, with added nanotopography, molded into PDMSe surface. ....................................... 40
2-6 AFM height contrast images of the same PDMSe nanotopography at different size scales. ......................................................................................................... 41
2-7 Average contact angle data of 5 µL droplets of DI water on Xiameter T2 surfaces .............................................................................................................. 41
2-8 The density of attached spores on PDMSe coatings after 45-minute settlement ........................................................................................................... 42
3-1 SEM micrograph of the fracture surface of silicon wafer with 250-2 topography etched into the top. .......................................................................... 53
3-2 SEM micrographs showing fracture surface of silicon wafers ............................. 54
3-3 Average RMS roughness data measured with AFM in tapping mode ................ 54
3-4 Average contact angle data of DI water on Bluesil surfaces ............................... 55
3-5 Average RMS roughness of 250-2 nanotopography on PDMSe surfaces after curing on PUR mold ........................................................................................... 55
3-6 The density of spores attached to nanopatterned PDMSe coatings after 45-minute settlement ............................................................................................... 56
3-7 The density of spores attached to nanopatterned PDMSe coatings after 45-minute settlement ............................................................................................... 56
4-1 AFM height contrast images of nanotopographies on various materials ............ 63
4-2 Average roughness of nanotopographies on various materials as measured by AFM in tapping mode ..................................................................................... 63
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4-3 RMS roughness data of final PDMSe sample topographies as measured by AFM in tapping mode. ........................................................................................ 64
4-4 Average contact angle of 5, 5 µL droplets of DI water on PDMSe surfaces. ...... 64
4-5 The density of spores attached to nanopatterned PDMSe coatings after 45-minute settlement ............................................................................................... 65
5-1 Reduced Young’s modulus as measured by AFM in PF QNM mode ................. 70
5-2 Adhesive force as measured by AFM in PF QNM mode .................................... 71
5-3 AFM images of silicone surfaces ........................................................................ 71
6-1 Inverted optical light microscope images of submerged microtopographies ....... 76
6-2 Inverted light microscope image of submerged +3SK2x2_n4 PDMSe topography after squirting surface with squirt bottle while submerged to induce wetting of the topography ........................................................................ 77
6-3 Inverted light microscope image of +8.5SK5x5_n6 PDMSe topography ............ 77
6-4 Inverted light microscope image of submerged +8.5SK5x5_n4_a8 PDMSe topography immediately after submersion in ASW ............................................. 78
6-5 Schematic representation of microtopographies................................................. 78
6-6 Inverted optical light microscope images of submerged PDMSe topographies after sonication in ASW ...................................................................................... 79
6-7 Inverted light microscope images of +8.5SK5x5_n9 PDMSe topography after conditioning and sonication ................................................................................ 79
6-8 Inverted light microscope images of +8.5SK5x5_n8 PDMSe topography after conditioning and sonication ................................................................................ 80
7-1 AFM height contrast images of SBS surface with no apparent organized nanoscale phase segregation ............................................................................. 87
7-2 AFM height contrast images of SBS surface after ozone removal of polybutadiene ..................................................................................................... 87
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LIST OF ABBREVIATIONS
AFM Atomic Force Microscopy
ATS Allyltrimethoxysilane
B. Amphitrite Balanus amphitrite
DMT Derjaguin, Muller, Toropov
DRIE Deep Reactive Ion Etching
ERI Engineered Roughness Index
HMDS Hexamethyldisilazane
ISK Inverse Sharklet
L-B Langmuir-Blodgett
PDMS Polydimethylsiloxane
PDMSe Polydimethylsiloxane elastomer
PEG Polyethylene glycol
PET Polyethylene terephthalate
PFOTS Trichloro (IH, IH, 2H, 2H – perfluorooctyl) silane
PF QNM PeakForce Quantitative Nanomechanical Mapping
PR Photo resist
PTFE Polytetrafluoroethylene
PUR Polyurethane
RIE Reactive Ion Etching
RMS Root mean square
SAM Self assembled monolayer
SEA Surface Energetics Attachment
SG184 Sylgard 184
SK Sharklet
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TBT Tributyltin
T2 Xiameter T2
U. linza Ulva linza
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Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy
EFFECT OF NANOSCALE AND HIERARCHICAL TOPOGRAPHIES ON THE
ANTIFOULING EFFICACY OF SILICONE SURFACES
By
Clayton Walker Argenbright
August 2018
Chair: Anthony Brennan Major: Materials Science and Engineering
Marine biofouling is a serious global issue which negatively impacts multiple
industries. The fouling of ship hulls decreases speed and fuel efficiency and transports
invasive species from one port to another. Current research is focused on nontoxic
alternatives to currently used toxic coatings which can be environmentally harmful.
Engineered microtopographies have shown promise but no one topography has been
shown to work against a variety of fouling species.
We have developed a process to add nanotopographies to microtopographies to
interact with organisms and structures on various size scales, and to change the wetting
behavior of the surface. Topographically modified surfaces were characterized primarily
using contact angle and Atomic Force Microscopy (AFM) to measure topography
dimensions and surface roughness.
A bioassay with Ulva linza zoospores showed that a nanotopography molded into
a polydimethylsiloxane elastomer (PDMSe) surface might reduce the settlement of algal
spores. Due to the discontinuation of this PDMSe by the manufacturer, the bioassay
could not be replicated on the same material. New PDMSe materials were acquired but
they did not replicate the nanotopographies well, and the reduction in spore settlement
14
was not seen in later assays. Differences in performance of smooth PDMSe surfaces
during testing show that more testing is required before definitive conclusions can be
drawn.
15
CHAPTER 1 INTRODUCTION
Scope of Research
Marine biofouling is a global issue with serious environmental and economic
consequences. This issue was previously handled using toxic coatings to kill fouling
organisms. Many of these coatings have recently been banned due to accumulation in
the environment and toxic effects on non-target organisms. These coatings need to be
replaced with nontoxic coatings that can still target a variety of fouling organisms. There
are many examples in nature that show that chemistry is not the only way to prevent
biofouling. For example, the skin of sharks and the shells of some mussels and crabs
show that surface topography can also be used as an effective antifouling strategy
(Bers & Wahl, 2004) (Greco, et al., 2013) (Kesel & Liedert, 2007) (Scardino, Hudleston,
Peng, Paul, & de Nys, 2009). By creating microscale, nanoscale, and dual-scale surface
topographies with regular, periodic dimensions and well-defined morphology, the effect
of topography on antifouling efficacy can be more accurately studied.
Specific Aims
Specific Aim 1: Develop method and fabricate periodic arrays of
nanotopographies and add them to existing microtopographies. Much of the work done
on nanoscale or hierarchical surfaces up to this point has been on very random
roughness. This is sometimes created by physically scratching the surface with an
abrasive material, or through chemical or plasma etching. In this study, both the
microscale and nanoscale topographies will be well defined and uniform over large
areas so that dimensions can be easily measured and systematically adjusted.
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Specific Aim 2: Determine the antifouling effects of nanoscale and dual-scale
surface topographies. The settlement of the zoospores of the green algae Ulva linza
has been used by the Brennan Research Group and other in the past to gauge the
antifouling potential of surfaces. That settlement assay will be used in this study as well
to compare results to previously tested PDMSe topographies.
Biofouling
Biofouling is the process of biological matter (proteins, cells, microorganisms,
and macroorganisms, etc.) attaching to and growing on a surface. This is a natural
process that occurs on most surfaces and progresses over time, often resulting in
complex environments consisting of multiple species that can be difficult to remove from
the surface. In the marine environment this typically occurs in 4 phases, which can
overlap chronologically, with the initial types of fouling molecules and organisms
continuing to foul the surface throughout its lifetime (Ecol Prog Ser & Wahl, 1989). The
first phase is the biochemical conditioning of the surface through the adsorption of
macromolecules. Polysaccharides, glycoproteins, and proteoglycans begin to
accumulate on the surface immediately upon immersion and can reach a dynamic
equilibrium within hours. The same types of molecules foul surfaces that begin with high
or low surface free energy such that the surface properties converge, often resulting in
surface free energy values of 30-40 mN/m (Baier R. , 1981). About an hour after
immersion, bacterial colonization of the surface begins through adsorption, then
adhesion of various bacteria. The growing bacterial colonies, their secretions, and the
underlining conditioning film are together known as a biofilm. Days after immersion,
unicellular organisms like protozoa, yeasts, and diatoms colonize the surface. Finally,
after days or weeks of immersion, multicellular organisms like algal spores and barnacle
17
cyprids begin attaching to and growing on the surface. Understanding and controlling
this process is of vital importance to various marine and medical industries.
In marine environments one of the most problematic areas where biofouling
occurs is on the hulls of ships and submarines where the accumulation of material
causes a variety of issues. As biofouling accumulates on a hull, the ship experiences
increased drag in the water, lowering fuel efficiency, acceleration, top speed, and
maximum range of the vessel. One full scale study found that after cleaning 22 months’
worth of microbial biofilm off the hull using brushes with relatively soft polypropylene
bristles, the ship required 18% less power to achieve a speed of 25 knots (Hasibeck &
Bohlander, 1992). The cleaning of this specific ship (the Knox class USS Brewton)
decreased fuel consumption by 350-600 gallons per hour, depending on speed. The
fouling penalty is even greater when the hull is fouled with larger organisms like algae
plants and barnacles. Just 5% coverage of a surface with shell fouling 14 mm in height
increases drag by 66%, and 75% coverage with shell fouling 4.5 mm in height would
increase the frictional resistance of a 120 m ship by 85% (Kempf, 1937). The increase
in drag leads to decreased fuel efficiency and speed, which increases the cost to
operate ships. Decreased speed of naval ships is also detrimental in a combat situation.
It is obviously beneficial to keep ship hulls free of fouling, but that is an expensive
prospect as well. It was estimated that hull fouling on just the DDG-51 destroyer class
ships costs $56 million per year (Schultz, Bendick, Holm, & Hertel, 2011). This cost
includes cleaning and recoating, but most of the cost comes from increased fuel
consumption. This figure represents just one of many classes of ships in the US Navy. If
that number is extrapolated out and applied to other naval and commercial shipping
18
vessels around the world, it is obvious the global economic costs of biofouling are
massive. Decreased fuel efficiency and speed from biofouling on vessels moving
consumer goods, food, oil or other raw materials contributes to the shipping cost of
those items.
Aside from the massive financial costs, biofouling can also have negative
environmental impacts. Transport of invasive species through biofouling is possible
because organisms can attach to a ship at one port, then be introduced at another port
half way around the world. One study found 34 unique multicellular fouling species on
the hulls of just 5 ships from the study that were analyzed (Davidson, Brown, Sytsma, &
Ruiz, 2009). Even the ships that had low overall accumulation of biofouling due to
relatively effective antifouling coatings still hosted a wide variety of species. They also
found that heterogeneous areas like recesses on the bottom of ships can reduce the
effectiveness of the antifouling coatings and host diverse colonies of fouling organisms.
Mussels growing in these areas can even provide a habitat for more motile organisms
that would otherwise not colonize the hull of a ship. Another study of international ships
in Osaka Bay, Japan, recorded 22 distinct species of barnacles on the hulls of just 2
bulk carriers. Fourteen of these species had never been recorded in Osaka Bay, and
another 2 were already considered invasive in Japanese waters. Many of these 14
species probably do not pose a threat of invasion in Osaka Bay specifically, due to
environmental factors, but at least 3 of the species were suitable to survive in that
environment and have already established invasive populations in other areas of Japan,
Europe, and South Africa (Otani, et al., 2007). This is a major concern as some invasive
19
species can completely alter an ecosystem and are extremely difficult to remove once
established, like the well-known example of the Zebra mussels in the Great Lakes.
Antifouling Strategies
Humans have been battling against biofouling for millennia. Ancient seafaring
cultures coated the hulls of ships with various types of pitch, tar and wax, sometimes
containing arsenic or sulfur to prevent shipworms from destroying wooden hulls (Callow
M. , 1990). There is also evidence of the ancient Phoenicians, Greeks, and Romans
using lead sheeting to protect wooden hulls, and the practice continued for thousands of
years until the middle ages (WHOI, 1952). Copper sheeting was used at least as early
as the 18th century, but copper was used in shipbuilding as nails and fasteners for
thousands of years previously (Borkow & Gabbay, 2009) and its antifouling properties
were certainly noticed.
After the development of iron ships, the use of external metal sheeting for
antifouling purposes was mostly abandoned due to increased galvanic corrosion
experienced by the hull and the excessive cost of insulating the hull from the metal
sheeting. Copper has remained in use as an antifouling material along with other toxic
metals like mercury and arsenic by the incorporation of biocidal compounds into
antifouling paints and coatings. The development of tributyltin (TBT) compounds in the
1950s was a significant step in antifouling technology (Dafforn, Lewis, & Johnston,
2011). TBT is a broad spectrum biocide that can remain effective for up to 5 years
depending on the type of coating it is incorporated into (Lewis, 1998). It was first used in
“free association” paints where it was dispersed within the paint and diffused to the
surface where it was desired. This type of coating resulted in unnecessarily high initial
release rates that would decrease and become ineffective relatively quickly as the
20
biocide was depleted from the coating. Eventually TBT was developed into a self-
polishing copolymer coating where the TBT was part of the paint matrix. As the biocide
dissolves into the surrounding sea water a fresh biocidal surface is constantly revealed.
The effective lifetime of these coatings is therefore proportional to the thickness of the
coating and can remain useful for years. Unfortunately, the effectiveness of both types
of these biocidal coatings rely on the release of a broad spectrum toxin into the
environment.
It became apparent in the 1980s that the accumulation of tin compounds in the
environment was having serious negative effects on the shell formation and
reproductive organs of various bivalves and gastropods (Alzieu, Sanjuan, Deltreil, &
Borel, 1986) (Omae, 2003) (Pangam, Giriyan, & Hawaldar, 2009) (Turner, 2010). It was
also estimated that the bioaccumulation of the toxins in fish and other organisms was
likely much higher than the levels measured in sea water and sediments (Omae, 2003).
These findings led to bans being gradually implemented in some areas on certain
vessels, until 2001 when the International Maritime Organization adopted rules to ban
the application of new TBT coatings by 2003 and recoat all ships with TBT free coatings
by 2008 (Dafforn, Lewis, & Johnston, 2011). Not all nations have agreed to these
regulations so there are still ships on the ocean using TBT based antifouling coatings.
Copper compounds and other biocidal alternatives for TBT are being used again as a
replacement, but the long term environmental impacts of these compounds is still
unknown. Current research in the industry is focused on developing non-toxic
antifouling fouling coatings, which release nothing into the environment and therefor
have no potential for toxicity, toxic degradation products, or bioaccumulation.
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Non-toxic Antifouling Strategies
A wide variety of approaches in this category are being developed and tested,
mostly involving altering the chemical or physical structure of a surface. One way to
alter the chemistry is by using self-assembled monolayers (SAMs) or polymeric grafts. It
has been found that hydrophilic surfaces, like those made from polyethylene glycol
(PEG) grafts and hydrogels, can reduce protein adsorption (Ostuni, Chapman, Holmlin,
Takayama, & Whitesides, 2001) (Ekblad, et al., 2008). When fouling proteins adsorb to
a submerged surface they must displace the water molecules from the surface.
Removal of water from around PEG chains is thermodynamically unfavorable due to the
decrease in conformational entropy upon dehydration. Longer polyethylene oxide (PEO)
chains grafted to a surface at high surface density have also been shown to decreased
protein adsorption through steric repulsion (Jeon, Lee, Andrade, & De Gennes, 1991). A
variety of hydrophobic surfaces have been tested as well. Polydimethylsiloxane
(PDMS) surfaces and fluorinated surfaces have shown some success in decreasing
settlement of, and facilitating the removal of the zoospores of the green algae Ulva
Linza and of barnacle cyprids, Balanus Amphitrite (Hu, et al., 2009) (Marabotti, et al.,
2009) (Martinelli, et al., 2011). These hydrophobic surfaces, however, allow proteins to
adsorb strongly to the surface through hydrophobic interactions and also do not
decrease the attachment of the diatom Navicula perminuta, another common marine
fouling organism (Krishnan, et al., 2006). For practical application, an antifouling surface
must be able to inhibit a broad range of organisms from attaching, not just one or two
species. This is a challenge since there are organisms that prefer to attach to
hydrophilic surfaces and those that prefer hydrophobic surfaces. To address this issue
chemically, people have experimented with surfaces that have zwitterionic or
22
amphiphilic character in order to repel multiple fouling species. A zwitterionic surface
with positively and negatively charged groups could potentially repel organisms which
prefer either charge state. It is the same idea with amphiphilic surfaces, organisms that
attach more easily to hydrophobic or hydrophilic surfaces may both have trouble
attaching to an amphiphilic surface (Carr, Xue, & Jiang, 2011) (Jiang & Cao, 2010)
(Krishnan, et al., 2006) (Park, et al., 2010) (Zhang, et al., 2009).
Antifouling Surface Topographies
Another natural antifouling strategy that shows promise is the use of physical
topographies on the surface. Many of these were inspired by fouling resistant surfaces
in nature which often have multiple scales of roughness. Microscale structures have
been shown to affect the ability of marine fouling organisms to attach to a surface. It
was found that the settlement density of the green alga Ulva linza in microscale
channels formed in a PDMS elastomer (PDMSe) is dependent on the size of the
channels in relation to the size of the organism. The Ulva spores are 5µm in diameter
on average, and settlement density was found to be the highest in channels that were
5µm wide and 5µm deep. It was later found that reducing the size and spacing of the
features to 2µm wide and 2µm apart (smaller than the size of an Ulva spore) greatly
reduced the settlement density of Ulva when compared to smooth PDMSe surfaces
(Callow, et al., 2002) (Carman, et al., 2006) (Schumacher, et al., 2007).
Wetting Behavior
The addition of topography to a surface changes the wetting behavior of that
surface, which may also contribute to antifouling potential. Wetting behavior on textured
or topographically patterned surfaces generally falls under one of two categories,
Wenzel or Cassie-Baxter style wetting. In the Wenzel state, the liquid completely fills the
23
pores or grooves in a surface. The spreading or receding of a drop of liquid on a solid
surface depends in part on the surface energy per unit area of the solid surface. A
roughened surface has more actual surface area than the flat projection of that surface,
so the same amount of surface energy is concentrated into a smaller projected area.
This causes roughness to enhance the wetting behavior of surfaces, making hydrophilic
surfaces more hydrophilic, and hydrophobic surfaces more hydrophobic (Wenzel,
1936). Wenzel used the roughness factor, r, to assign a value to rough surfaces.
Wenzel’s roughness factor is defined in Equation 1-1.
𝑟 = 𝑟𝑜𝑢𝑔ℎ𝑛𝑒𝑠𝑠 𝑓𝑎𝑐𝑡𝑜𝑟 =𝑎𝑐𝑡𝑢𝑎𝑙 𝑠𝑢𝑟𝑓𝑎𝑐𝑒 𝑎𝑟𝑒𝑎
2𝐷 𝑝𝑟𝑜𝑗𝑒𝑐𝑡𝑒𝑑 𝑠𝑢𝑟𝑓𝑎𝑐𝑒 𝑎𝑟𝑒𝑎 (1-1)
The apparent contact angle on a rough surface (θ*) can then be related to the
contact angle from Young’s equation, θ, in Equation 1-2 on a homogeneous surface by
Equation 1-3.
𝛾𝑆𝑉 = 𝛾𝑆𝐿 + 𝛾𝐿𝑉 cos 𝜃 (1-2)
cos 𝜃∗ = 𝑟 cos 𝜃 (1-3)
In the Cassie-Baxter regime, air pockets remain trapped under the liquid in the
roughness of the surface. The result is a composite interface where the liquid sits on
solid and vapor, rather than a homogeneous interface as seen in the Wenzel state
(Cassie & Baxter, 1944). The resulting equation taking this composite interface into
account is Equation 1-4.
cos 𝜃∗ = 𝑓1 cos 𝜃 − 𝑓2 (1-4)
In Equation 1-4, 𝑓1 is the area of solid-liquid interface and 𝑓2 is the area of liquid-
vapor interface (in the same plane as the solid-liquid interface). Decreasing the contact
24
area with the solid surface in this way leads to increased contact angle, and is often
seen in hydrophobic and superhydrophobic surfaces.
Wettability of a surface can also influence the fouling behavior of the surface.
Researchers have tried to correlate surface energy to fouling behavior of a surface, with
the most notable trend being represented with the Baier Curve (Baier & DePalma,
1971). This model predicts that surfaces with critical surface tension between 20 and 30
mN/m will be the least susceptible to fouling. However, the variety of fouling organisms
found in nature limit the effectiveness of these models in predicting fouling behavior
because different organisms have been found to preferentially settle on surfaces with
different energies (in hydrophilic and hydrophobic regimes). Also, the surface on which
the organism attaches is often chemically and physically different from the original
surface due to conditioning layers of proteins and minerals deposited through contact
with the liquid. The Brennan Research Group and collaborators have demonstrated this
variability of fouling organisms and behaviors by testing a wide variety of surface
topographies and chemistries against multiple types of organisms (Hoipkemeier-Wilson,
et al., 2004) (Holland, et al., 2004) (Schumacher, et al., 2007) (Schumacher, et al.,
2007). The Cassie-Baxter wetting state and superhydrophobicity may offer a way
around these problems. By limiting the contact area of the liquid with the surface you
are limiting the amount of surface area which could potentially become contaminated.
Consider the extreme case of a perfectly hydrophobic surface on which a water drop
has a contact angle close to 180°. Since the water is not wetting the surface, any solute
(proteins, minerals, etc.) or microorganisms in the water will not contact the surface, and
therefore will not attach to it.
25
Conclusion
Biofouling is a complicated and dynamic process which causes a lot of harm and
needs to be addressed. There are multiple avenues currently being researched to
develop nontoxic antifouling coating which can work against a variety of fouling
organisms. This work will continue previous work of the Brennan Research Group by
focusing on surface topographies to physically prevent the settlement of organisms. It
will expand on the previous work by adding a smaller scale of topography to existing
microtopographies, altering the wetting properties of the surface and hopefully the
interaction with multiple fouling organisms.
26
CHAPTER 2 ADDITION OF NANOTOPOGRAPHY TO ANTIFOULING SILICONE
MICROTOPOGRAPHY
As discussed in the previous chapter, marine biofouling is a major issue with
global economic and environmental consequences. Previous efforts to combat this
problem on ships often relied on toxic hull coating, which have caused their own
environmental issues like toxicity to non-target organisms. The Brennan Research
Group has been investigating microscale surface topographies as a physical means to
prevent marine biofouling on a surface. The Sharklet AF topography has been
successful at reducing the settlement of zoospores of the green algae Ulva linza,
potentially due to the size of the topography relative to the size of the spores. This effect
is not seen with organisms of other sizes on this same size microtopography. The
addition of nanotopography to the microtopography could allow for the surface to
interact with organisms of other sizes and could change the wetting behavior of the
surface by increasing roughness and moving toward a more extreme wetting or non-
wetting state.
Background
Microtopographies
Microscale surface topographies are prevalent in nature where they give plants
and animals antifouling properties due to unique non-wetting behaviors. A common
example is that of the Lotus leaf which has microscale bumps, coated with a
hydrophobic wax which also provides nanoscale roughness (Koch, Bhushan, &
Barthlott, 2009). This structure gives the lotus leaves Superhydrophobic and self-
cleaning properties. Lesser known examples include the bumpy microscale structures
on the shells of Cancer pagurus crabs and the surface of Ophiura texturata brittle stars
27
which have been shown to reduce the settlement of various marine fouling species
(Bers & Wahl, 2004). Many mollusk shells have naturally occurring microstructures and
antifouling properties as well, such as the periodic ribbed structure of Dosinia juvenilis,
and the periodic spines on the shells of Tellina inflata (Scardino, Hudleston, Peng, Paul,
& de Nys, 2009).
Microtopographies have been shown to be able to both enhance and reduce
settlement of some organisms based on the size of the organisms and the surface
features. For example, the spore of the green algae Ulva linza is approximately 5
microns in diameter. It was found that in grooves molded into PDMSe surfaces with
width and spacing of 5 microns, spores could fill in the channels and attachment density
increased compared to smooth PDMSe surfaces (Callow, et al., 2002). If the width and
spacing of topography was reduced to 2 microns like in the case of the Sharklet AF
microtopography, the spores could not squeeze down into the channels and settlement
was reduced compared to smooth PDMSe (Carman, et al., 2006) (Schumacher, et al.,
2007). A similar size relationship was seen with the larger 20 µm x 20 µm Sharklet AF
topography which was able to reduce the settlement of the larger Balanus amphitrite
cyprids (barnacle larvae) (Schumacher, et al., 2007). Currently, the best model to
explain and predict this behavior for various organisms is the Surface Energetics
Attachment (SEA) model developed by Decker et al. shown in Equation 2-1 (Decker, et
al., 2013), where Nt and Ns are the number of organisms attached to the topography
and smooth surfaces respectively, A is the interfacial area between the organisms and
the surface, and g is the number of available settlement sites on the surface.
ln (𝑁𝑡
𝑁𝑠) =
⟨𝐴𝑡⟩−𝐴𝑠
𝐴𝑠+ ln (
𝑔𝑡
𝑔𝑠) (2-1)
28
The SEA model is partially based on the attachment point theory, which is based
on the observation that fouling organisms tend to attach where they can make the most
number of contacts with the surface, resulting in more stable attachment. A good
example of this is the increase in U. linza settlement in the 5 µm wide, 5 µm deep
channels as seen by Callow et al. and discussed above. This theory predicts fouling
behavior well for some organisms, but not at all for others, especially non-motile fouling
organisms which do not “choose” where to settle (Scardino, Guenther, & de Nys, 2008).
The SEA model also takes into account the contact area that is actually available to the
organism due to the roughness of the surface and the potential for Cassie-Baxter style
non-wetting behavior. This model is the first to accurately predict both the inhibition and
enhancement of the attachment of miltiple different fouling organisms to a topography.
Hierarchical Topographies
By adding a second size scale of topography on the microtopography the surface
may be able to interact with organisms of multiple size scales. As previously discussed,
the relationship between the size of features on a topography and the size of an
organism, or some part of that organism, is important in determining if that topography
can interact meaningfully with that organisms. If one is much larger or smaller than the
other, the organisms may simply sense a smooth surface. The addition of
nanotopography to microtopography could enhance the interaction of the surface with
smaller cells, like bacterium, or with specific cell membrane structures on larger cells.
As discussed in the previous chapter, aside from direct physical interaction with
organisms, the second scale of topography can also change the wetting behavior of the
surface. On a hydrophobic surface like PDMSe, the addition of roughness increases the
surface area and therefore enhances the hydrophobic behavior. The Sharklet AF
29
microtopography shows a metastable wetting state when submerged in water, holding
air in the channels for a period of time before eventually filling with water. The addition
of another scale of topography to increase roughness and surface area could make the
surface harder to wet and help it maintain the Cassie-Baxter non-wetting state for
longer.
Objectives
Objective 1: Add nanotopography to microtopography to create hierarchical
surface topography in PDMSe surfaces. The hierarchical topography will be able to
interact with organisms of more size scales that the microtopography alone. It will also
make the hydrophobic PDMSe less wettable by increasing the roughness and therefore
the hydrophobic surface area, which could help delay fouling in a marine environment.
Objective 2: Evaluate antifouling potential of nanopatterned and hierarchical
topographies in comparison to previously tested microtopographies. Antifouling potential
will be evaluated by measuring the settlement density of Ulva linza zoospores on the
topography after the assay. This test has been used frequently in the past and has
showed the effectiveness of the Sharklet AF microtopography. In this study the
microtopography, nanotopography, hierarchical topography, and smooth PDMSe will all
be tested for comparison.
Materials
Single crystal silicon wafers (100 mm diameter, <100> orientation, prime grade)
were purchased from University Wafer. Hexamethyldisilazane (HMDS),
allyltrimethoxysilane (ATS) and ethylene glycol were purchased from Sigma Aldrich.
Shipley AZ1512 photoresist (PR), AZ 300 MIF developer, and PRS3000 (PR stripping
solvent) are purchased and supplied by the Nanoscale Research Facility at the
30
University of Florida. Spherical silica nanoparticles (250 nm diameter) were purchased
from Particle Solutions, LLC. DI water was made using a Thermo Scientific Barnstead
Nanopure water system. Platinum catalyzed polydimethylsiloxane elastomer (PDMSe)
(Xiameter T2, made by Dow Corning) was purchased from Essex-Brownell Inc.
Thermoplastic polyurethane pellets (PUR) were purchased from Lubrizol Advanced
Materials, Inc. Ethanol and acetone were purchased from Fisher Scientific. Ultra high
purity nitrogen gas was purchased from Airgas.
Methods
Microscale Patterning
Silicon wafers were opened in the clean room, then treated with vapor deposition
of HMDS on a 112°C hot plate. Wafers were then coated with PR using a Suss Delta 80
spin coater (0.8 µm thick). Microscale patterns (SK2x2, InvSK2x2) were drawn using
LayoutEditor software, then directly written into the photoresist using a Heidelberg DWL
66FS Maskless Laser Lithography System (DWL). The parameters of the DWL for
optimal exposure vary over time with laser power and focus. Dosage was optimized
each time by writing and developing a test grid on a dummy wafer. After development of
the photoresist the wafers were etched in an STS DRIE. Two-micron wide patterns were
etched using a continuous plasma etching process where SF6 and C4F8 gasses are
used simultaneously. SF6 etches away the exposed silicon while C4F8 deposits on the
surface and acts as a passivation layer to prevent etching. When used simultaneously
the result is a microscale topography with smooth side walls, when cycled the sidewalls
have small scallops but higher aspect ratio etching is possible. PR was then removed
from the wafer by soaking in PRS3000 at 70°C for 15 minutes. These procedures were
used to make both the Sharklet AF (SK) microtopography, as well as the Inverse
31
Sharklet AF (ISK) microtopography (in which the rectangular features are depressed
holes instead of raised ridges).
Langmuir-Blodgett Coating
Smooth wafers were treated with air plasma at 500 mTorr, 30 Watts, for 1 minute
in a Harrick Plasma Cleaner. The wafers were then coated with silica nanoparticles
using the Langmuir-Blodgett (L-B) method (Yang, Dou, Fang, & Jiang, 2013). The
wafers were secured (one at a time) to a large syringe pump, oriented to move
vertically, and submerged in a 110 mm wide, 1000 mL beaker of DI water. A dispersion
of nanoparticles in ethylene glycol (~1.5 wt.%) was slowly and carefully dripped around
the inside edge of the beaker using a pipette until a close packed layer of particles was
formed floating on the surface of the liquid. The syringe pump was then turned on,
drawing the wafer up out of the liquid (with the long axis of the microtopography pointing
upward), at a rate of about 9 mm/min while more nanoparticle dispersion was
continually added around the edge. The result is a close packed monolayer (with
defects) of silica nanoparticles deposited on the surface of the wafer.
Reactive Ion Etching
The 250 nm particle coated wafers were then etched in a Unaxis Shuttlelock
Reactive Ion Etcher (RIE) with SF6 plasma for 30 s (RF power at 150 Watts, 25 °C, 10
mTorr, 50 sccm of SF6). The particles act as a mask during the isotropic etch, resulting
in a conical pillar left behind under each particle. After etching, the wafer is submerged
in a solution of 5% HF in water for 30 seconds, and then etched again in the RIE using
the same parameters for 5 seconds. The purpose of doing a second etch after the
particle mask has been removed is decrease the diameter of the pillars and increase
the height without creating an undercut, mushroom shaped morphology.
32
HMDS Treatment
Wafers were first rinsed with acetone, then ethanol, and dried with nitrogen gas.
They were then placed in an empty glass vacuum desiccator, vacuum was pulled for 10
min to a magnitude of approximately 150 mTorr. The desiccator was then closed to the
pump and opened to a bottle of HMDS via a syringe line for 10 minutes. The desiccator
was then sealed and placed in a preheated oven at 80 °C for 30 min.
Polymer Replication
Wafers were rinsed again with acetone, then ethanol, and dried with nitrogen.
PDMSe was mixed at a ratio of 10 parts base to 1 part curing agent and degassed
under vacuum. It was then poured on the wafers and sandwiched between two PET
sheets and two glass plates, where it cured for 24 hours. This PDMSe curing process
was done twice on each wafer before attempting PUR replication. PUR pellets were
treated in an oven at 120°C for 2 hours. Approximately 5 grams of pellets were placed
in a square metal mold 1 mm thick, sandwiched between two layers of PTFE film and
two steel plates. The whole assembly was placed in a Carver heated hydraulic press at
180 °C for 3 minutes, then pressed with a pressure of 2 metric tons into a flat film about
2 mm thick. A piece of this PUR film large enough to cover the micropatterned area of
the wafer (at least 25 mm x 25 mm) was cut and placed on the pattern on the wafer.
The wafer was then placed between the PTFE and steel plates, the film was melted
again and pressed into the topography of the wafer using the same temperature and
pressure as the first melt. The wafer was removed from the press and placed on a lab
bench to briefly cool, the PUR was peeled off the wafer as soon as it was cooled
enough to release cleanly from the wafer (determined by peeling up the edge of the
33
PUR). The patterned PUR was then replicated with PDMSe using the same procedures
described above, with no additional surface treatment.
Sample Mounting
PDMSe sample films were trimmed into 25 mm by 25 mm squares. They were
rinsed with acetone, then ethanol, and dried with nitrogen. They were placed
topography side down on clean, smooth, HMDS treated glass plates. Glass microscope
slides were flame treated with a propane torch, then coated with ATS using a solution
deposition process in ethanol. The ethanol and ATS solution was rinsed off with ethanol
and slides were dried in an oven at 120°C for 10 minutes, and allowed to cool at room
temperature. A fresh batch of PDMSe was mixed, degassed, and a thin layer was
poured over the backs of the previously cured PDMSe sample films. ATS treated slides
were secured to another HMDS treated glass plate using double sided tape, the two
glass plates were then pressed together against spacers such that each PDMSe
sample film ends up in the center of a glass slide. After the new PDMSe was cured, the
plates were separated, and the microscope slides were cut out of the PDMSe sheet.
The result was a microscope slide completely covered in PDMSe of uniform thickness
with the sample topography mounted in the center and surrounded by smooth PDMSe.
Characterization
Characterization of wafer and polymer surfaces was performed after each
processing step by measuring contact angles on a Rame-Hart goniometer, and imaging
surface topography with a Bruker Dimension Icon AFM with ScanAsyst, and an FEI
Nova 430 SEM. PDMSe surfaces were sputter coated with Gold/palladium in a Denton
Vacuum Desk II sputter coater prior to SEM imaging. Due to the asymmetry of the
34
Sharklet AF microtopography, contact angles were recorded and reported by viewing
droplets parallel to and perpendicular to the long axis of the topography.
Ulva linza Settlement Assay
Ulva assays were conducted by John Finlay et al. at Newcastle University
according to their established procedures. Coatings were equilibrated in 0.22 μm filtered
artificial seawater for 24 hours prior to testing. Zoospores were obtained from mature
plants of U. linza by the standard method. A suspension of zoospores (10 ml; 1x106
spores ml-1) was added to individual compartments of quadriPERM dishes containing
the samples. After 45 minutes in darkness at 20 °C, the slides were washed by
immersion in fresh artificial seawater to remove unsettled (i.e. swimming) spores. Slides
were fixed using 2.5% glutaraldehyde in seawater. The density of zoospores attached to
the surface was counted on each of 3 replicate slides using an image analysis system
attached to a fluorescence microscope. Spores were visualized by autofluorescence of
chlorophyll. Counts were made for 30 fields of view (each 0.15 mm2) on each patterned
area.
Results and Discussion
Wafer Mold Fabrication
The L-B coating technique resulted in a close packed monolayer with defects
(Figure 2-1). Defects include grain boundaries, holes, and a few particles sitting on top
of the monolayer. Some defects are expected as there is some particle size variation.
The coating was consistent across the entire coated area of the wafer. After etching in
the RIE, a pillar remains below the center of each particle. These pillars are about 130
nm tall and have flat, roughly circular tops approximately 100 nm in diameter (Figure 2-
2)(Figure 2-3). The hexagonal packing order of the particle monolayer remains for the
35
pillars, and the topography (with some defects etched in) is consistent across the wafer
surface.
Polymer Replication
HMDS treatment increased the contact angle by over 40° on the nanotopography
(Figure 2-4), and successfully prevented bonding of PDMSe to the wafer surface. After
two replications with PDMSe, the PUR mold was pressed on the wafer and peeled off
cleanly without leaving residue behind. PDMSe was then cured on the PUR with no
further surface treatment. The resulting PDMSe surface has the nanotopography
molded onto the top of the microtopography as well as on the bottom of the channels
(Figure 2-5). The intention was to create the standard +2.6SK2x2 microtopography
(meaning Sharklet AF topography which is 2 microns tall, 2 microns wide, spaced 2
microns apart), however after the extra etching steps and multiple replications the final
dimensions were actually +1.5SK1.2x2.4. The average contact angles on the
hierarchical topography are 129.8° ± 3.2° (perpendicular) and 142.7° ± 3.3° (parallel),
compared to 132.9° ± 2.4° (perpendicular) and 139.0° ± 3.0° (parallel) on the standard
+2.6SK2x2 microtopography with no nanotopography added. These values indicate that
the droplet on the hierarchical topography is still sitting on top of the microtopography in
the Cassie-Baxter non-wetting state, but may be wetting the nanotopography in the
Wenzel state. It should be noted that deposition of the droplet on the hierarchical
topography was more difficult than on the microtopography, most attempts resulted in
the droplet remaining stuck on the end of the syringe, and the hierarchically patterned
surface remaining dry. Multiple attempts were required to deposit the droplets on the
surface which means that the measurements may have been made on areas that were
less hydrophobic, less rough, or where defects or contamination was present. This
36
could skew the average contact angle to a lower value. The PDMSe nanotopography
had an RMS roughness of about 24 nm as measured by AFM. This value varies slightly
depending on scan parameters during AFM imaging. For example, Figure 2-6A has
RMS roughness of 24.5 nm while Figure 2-6B has RMS roughness of 23.5 nm. In both
cases the measured increase in surface area (compared to a projected smooth surface
of the same planar area) was greater than 10%. This was enough to increase the
average contact angle from about 109° to almost 117° (Figure 2-7). This is slightly lower
than the expected apparent contact angle based on the Wenzel equation at this level of
roughness. A more recent analysis of contact angle measurements has led to a
modification in this equation to show that the contact angle of a drop on a surface is not
dependent on the roughness below the entire drop, but rather the roughness at the
contact line (Seo, Kim, & Kim, 2015). A drop of water on a solid surface (especially a
soft one) can deform the surface at the contact line as well (Hui & Jagota, 2014)
(Jerison, Xu, Wilen, & Dufresne, 2011) (Leh, et al., 2012) (Style, Che, Wettlaufer, Wilen,
& Dufresne, 2013), in this case possibly leading to lower roughness and contact angle.
These contact angle values show that the protrusion of the nanotopography are not tall
enough to support a drop in the Cassie-Baxter non-wetting state, the drop instead wets
the nanotopography in the Wenzel state.
Ulva linza Settlement Assay
The spore settlement density was significantly lower on all topographies than on
the smooth PDMSe (Figure 2-8). The ISK had the highest settlement density of the
patterned films, this was due to 1 of the 3 replicates tested seeming to be an outlier.
Two of the three ISK samples performed similarly to the other topographies. The
addition of the nanotopography to the SK and ISK microtopographies did not further
37
decrease the settlement density, they instead performed similarly. The most interesting
result of this study was that the nanotopography alone (no microtopography) performed
as well as any of the other topographies tested. The addition of the nanotopography to
smooth PDMSe reduced the settlement density of spores by 74%.
Conclusion
Periodic, close packed nanotopography was successfully added to smooth and
micropatterned silicon wafers. The topographies were successfully replicated in PUR
and PDMSe. The addition of the nanotopography to the Sharklet AF microtopography
did not appear to increase or decrease the antifouling efficiency of the surface. The
nanotopography on otherwise smooth PDMSe performed as well as any of the other
topographies. This result is very interesting because nanotopography of this type can be
added to a micropatterned wafer much more easily than a second microtopography
could be added. If algae spore settlement can be reduced by just the nanotopography, it
could potentially be added to a larger microtopography designed to deter larger
organisms, like barnacle cyprids. The result would be a physical topography with the
ability to deter multiple organisms of varied sizes. The next step is to confirm these
results with further testing and expand the size range of nanotopographies tested to
optimize the nanotopography before adding it to a new microtopography.
38
Figure 2-1. Two dimensional and three dimensional representations of silica particle
monolayer on wafer surface imaged with AFM in ScanAsyst mode.
Figure 2-2. SEM micrograph of the fracture surface of a silicon wafer with nanotopography etched into the surface.
39
Figure 2-3. SEM micrographs of nanotopography etched into silicon wafer surface. The
surface was tilted at a 45° angle relative to the electron beam for imaging. A-D are images of the same surface at different size scales. D) A hexagonal overlay has been added to the image to highlight the hexagonal close packed pattern of the topography which was transferred from the silica particle monolayer.
40
Figure 2-4. Average contact angle data from water droplets on wafer and PDMSe
surfaces throughout replication process. Error bars represent one standard deviation above and below the mean.
Figure 2-5. SEM micrograph of part of a unit cell of Sharklet AF microtopography, with
added nanotopography, molded into PDMSe surface.
41
Figure 2-6. AFM height contrast images of the same PDMSe nanotopography at different size scales. Both images were collected from the same surface using ScanAsyst mode.
Figure 2-7. Average contact angle data of 5 µL droplets of DI water on Xiameter T2
surfaces, error bars represent one standard deviation above and below the average.
42
Figure 2-8. The density of attached spores on PDMSe coatings after 45-minute
settlement. Each point is the mean from 90 counts from 3 replicate slides. Bars show 95% confidence limits
43
CHAPTER 3 EFFECT OF POLYDIMETHYLSILOXANE NANOTOPOGRAPHIES ON THE
SETTLEMENT OF THE SPORES OF Ulva linza ALGAE
Marine biofouling is a major economic, environmental and national security issue.
The buildup of biological material on the hulls of ships slows them down (Townsin,
2003), decreases fuel efficiency, and can transport invasive species around the world
(Otani, et al., 2007) (Piola, Dafforn, & Johnston, 2009). The U.S. Navy spends over $50
million annually on fouling related costs on just the DDG-51 class destroyers (Schultz,
Bendick, Holm, & Hertel, 2011). The process begins small with the adhesion of
biological molecules, algae spores, and other microorganisms to the surface creating a
biofilm. Larger fouling species like barnacle cyprids and tubeworm larvae then colonize
the surface as well. Toxic paints have often been used to deal with this problem but
have recently been banned due to the buildup in the environment and the toxicity to
non-target species. It is vital that new antifouling strategies are developed which are
non-toxic and effective against a variety of fouling organisms.
Background
Nontoxic Antifouling Strategies
A variety of antifouling strategies have been tested recently with mixed success.
Many surfaces have been created that can reduce the settlement of, or weaken the
attachment of a specific organism. Unfortunately, there is a wide variety of fouling
organisms in the world’s oceans, so an antifouling surface must be able to repel more
than one of them. For example, some organisms can attach strongly to hydrophilic
surfaces, and others to hydrophobic, so a single surface chemistry may not be effective.
Amphiphilic surfaces have been created by combining low surface energy polymers with
antifouling properties and hydrophilic chains like PEG which can resist protein
44
adsorption. These surfaces have been shown to be able to reduce settlement or ease
the removal of multiple fouling species and proteins (Krishnan, et al., 2006) (Martinelli,
et al., 2011) (Park, et al., 2010) (Weinman, et al., 2010). Mixing the right surface
chemistries and surface roughness can create extreme wetting states which could
potentially repel organisms as well. If a superhydrophillic surface can keep water bound
tightly, other organisms and molecules will have trouble displacing that water to form a
stable bond. Superhydrophobic surfaces can keep a layer of air trapped at the surface,
inhibiting organisms or molecules of water from even contacting the surface.
Unfortunately, these non-wetting states have proven to be metastable, and as wetting
occurs, the surfaces begin to foul (Marmur, 2006).
Nanotopographies
With recent technological advancements the ability to characterize and fabricate
nanotopographical surfaces has become simpler, cheaper, and more accessible. Like
the microtopographies discussed previously, nanotopographies are prevalent
throughout nature on the surfaces and internal structures of plants and animals and
have been shown to influence cells and organisms. It has been demonstrated that the
differentiation of human neural stem cells could be enhanced and directed by
nanotopographies with dimensions of 300-600 nm. The smallest patterns included in the
study had width and spacing of 300 nm and showed the most enhancement to
differentiation. The differentiation to either astrocyte or neuronal lineages was
determined by the geometry of the topographies, which were either square arrays of
square pillars or periodic channels (Yang, et al., 2013).
Nanotopographies have also been shown to have biocidal properties. The wings
of the Orthetrum villosovittatum dragonfly have cylindrical pillars of two sizes on their
45
surface. The small pillars are approximately 189 nm tall and 37 nm wide while the larger
pillars are about 311 nm tall and 57 nm wide. Escherichia coli bacteria that attach to this
surface die when their membranes are ruptured as they try to move to a more favorable
surface (Bandara, et al., 2017). Similarly, the wings of cicada also have
nanotopographies and biocidal properties at this size scale. Megapomponia intermedia
and Cryptotympana aguila cicada wings have hexagonally packed bumps on their
surface with height of 241 nm and 182 nm respectively, and pitch of 165 nm and 187
nm. Pseudomonas fluorescens bacteria were found to die on this surface as well
(Kelleher, et al., 2016). In a separate study on cicada wings, it was found that the
biocidal effect did not extend to more rigid bacteria. They theorized that as the less rigid
membranes conform to the topography, the area between features stretches and
ruptures (Pogodin, et al., 2013). The nanotopographies created in this study are similar
in geometry and scale to those found on the cicada wings.
Marine fouling and fouling removal properties are also influenced by
nanotopographies. The Carcinus maenas crab is able to keep its eyes extremely clean
using soft brush-like appendages without damaging the eye surface. Characterization
revealed deep, narrow, microscale grooves, surrounding flatter hexagonal regions
covered in nanotopography. The nanotopography was found to be consistent across
crabs tested in all stages of life, and had RMS roughness of about 16-20 nm (Greco, et
al., 2013). As previously discussed, settlement of spores of the U. linza algae is reduced
on topographies with dimensions around 2 µm. Cao et al. tested U. linza settlement on
more random topographies with dimensions down to 600 nm, and found that while
settlement increased with decreasing feature size, fouling removal was enhanced (Cao,
46
et al., 2010). In the previous chapter, a study of hierarchically patterned PDMSe
indicated that a hexagonally packed array of nanobumps (250 nm pitch) may be
effective at reducing settlement of the spores of U. linza (Figure 2-8). This study seeks
to expand the range of nanotopographies tested with this same morphology to pitches
of 100-500 nm for use in future nano- or hierarchically patterned antifouling surfaces.
Objectives
Objective 1: Fabricate series of nanotopographies in Bluesil RTV 3040 silicone
with dimensions from 100 nm to 500 nm. The previous study on hierarchical and
nanoscale topographies showed that the nanotopography alone had antifouling
potential (Figure 2-8). That same topography (250-2) will be included in this study to
replicate results, as well as a new 250 nm topography, and topographies created with
100 nm and 500 nm particle masks.
Objective 2: Evaluate settlement response of zoospores of U. linza green algae.
Chapter 2 showed that while the nanotopography may have had an antifouling effect, it
did not increase the antifouling efficacy of the Sharklet AF microtopography when added
to the top to make a hierarchical topography. By evaluating the effects of a larger range
of nanotopographies we can hopefully learn if larger or smaller nanotopographies work
better than the 250-2 topography. If the nanotopography can be optimized to prevent
settlement of zoospores, it could potentially be added to a larger microtopography
designed to deter another species, like barnacle larva, to make a broader range
antifouling surface.
Materials
Spherical Silica nanoparticles (100 nm, 250 nm, and 500 nm diameter) were
purchased from Particle Solutions, LLC. Bluestar Bluesil RTV 3040 (PDMSe) was
47
obtained from Bluestar Silicones. All other materials were obtained from the same
sources as in Chapter 2.
Methods
Langmuir-Blodgett Coating
Procedures for coating with 500 nm and 250 nm particles were the same as
described in Chapter 2. Procedures were slightly adjusted to handle the 100 nm
particles, an 8” wide crystallization dish was used for the water bath and the wafer was
removed from the bath at a slower rate of about 4 mm/min.
Reactive Ion Etching
The particle coated wafers were then etched in a Unaxis Shuttlelock Reactive Ion
Etcher with SF6 plasma using the parameters listed in Table 3-1. After etching once, the
wafers were submerged in a solution of 5% HF in water for 30 seconds to remove the
silica particles. Some samples were then etched again without a mask to attain the
desired morphology as described in Chapter 2. Samples made for this study (conical
morphology of wafer topography) are called 100-1, 250-1, and 500-1. The topography
tested previously and included again in this study will be called 250-2. Parameters not
listed in Table 1 were held constant for all etches (25 °C, 10 mTorr, 50 sccm of SF6).
Polymer Replication
Wafers were first rinsed with acetone, then ethanol, and dried with nitrogen gas.
Wafers and an open vial containing 0.1 mL of PFOTS were placed in vacuum
desiccator, vacuum was pulled for 5 minutes to a magnitude of approximately 180
mTorr. The desiccator was left in the fume hood sealed at room temperature for 1 hour.
It was then vented; the wafers were cleaned with acetone and ethanol again and dried
with nitrogen. They were then replicated with PDMSe twice, then with PUR using the
48
procedures described in Chapter 2. Samples were cut and mounted on microscope
slides according to the procedures in Chapter 2.
Characterization
Characterization of wafer and polymer surfaces was performed using the same
instruments and methods as described in Chapter 2. Roughness was measured by
AFM in tapping mode using Bruker MPP-11120-10 RTESPA tips. RMS roughness was
calculated by NanoScope Analysis 1.5 software using Equation 3-1 where Zi represents
the current Z value and N represents the number of points in the image.
𝑅𝑀𝑆 𝑅𝑜𝑢𝑔ℎ𝑛𝑒𝑠𝑠 = √∑(𝑍𝑖)2
𝑁 (3-1)
Ulva linza Settlement Assays
Ulva assays were conducted by John Finlay et al. at Newcastle University
according to their established procedures which were described in Chapter 2. Assays
were performed on two different sets of samples with the same topographies,
approximately 3 months apart. In Assay 1 the smooth areas surrounding the pattern,
and slides that were completely smooth (cured on HMDS coated glass) were used as
controls for comparison to patterned areas. In Assay 2, smooth samples were included
that were cured on PUR and HMDS coated silicon wafers, then cut and mounted using
the same procedures as for patterned films to ensure that they were processed as
similarly as possible to the patterned samples.
Results and Discussion
Nanotopographies
The 250-2 topography (Figure 3-1) was created during a preliminary study which
indicated the nanotopography alone had antifouling potential. Based on those results,
49
the new series of nanotopographies was created with different etching parameters. The
new parameters used to etch 100-1, 250-1, and 500-1 (Table 3-1) were developed to
increase the feature height and decrease the area of the feature tops. This difference
can be seen on the different 250 nm topographies shown in Figure 3-1 and Figure 3-2B.
This was intended to increase roughness on final PDMSe samples. Due to the spherical
morphology of the silica particles composing the hexagonally packed etching mask, the
etching rate varies under different areas of the particle during RIE. The areas of the
wafer directly below the holes where 3 particles meet etch the fastest, the areas below
where two particles touch etch slower, and the areas below the center of a particle etch
the slowest. In the etching processes used in this study, the original wafer surface that
is in contact with the particles is not etched away and remains on the final features such
that they all share the same top surface plane (with defects). Since feature height may
vary depending on where around the feature it is measured, maximum feature heights
will be discussed.
All topographies etching into silicon wafers consisted of conical pillars with flat
tops, roughly circular in shape, hexagonal packing order and pitch corresponding to
diameter of particles in mask (100, 250, or 500 nm). After etching using the parameters
in Table 3-1 the 500-1 topography had maximum feature height of 520 nm, and average
feature top diameter of ~230 nm. The 250-1 topography had maximum feature height of
190 nm with feature tops ~60 nm wide (compared to 130 nm tall and ~80 nm wide on
the 250-2 topography). The 100-1 topography had maximum feature height of 90 nm
and relatively sharp feature tops with diameters less than 10 nm.
50
Polymer Replication
The PUR hot embossing process did not work as well during this study as it did
in Chapter 2 when Xiameter T2 PDMSe was used. The PDMSe had to be replaced due
to the discontinuation of T2 by the manufacturer. The replication of the wafer with T2
prior to PUR embossing was a necessary step of the process to ensure the PUR
released cleanly from the wafer surface. After the switch to Bluesil RTV 3040 this
process did not work as well, when the PUR films were removed from the wafer surface
residue was left behind in some areas, leaving microscopic, random roughness on the
PUR surface. Procedures were adjusted as described above to maximize the area of
the PUR that released cleanly from the wafer, and care was taken to cut PDMSe
samples from only these areas which replicated well.
Unfortunately, surface roughness decreased multiple times throughout the
replication process (Figure 3-3), resulting in silicone surfaces much smoother than the
original silicon wafers shown in Figure 3-2. After hot embossing of the wafer with PUR,
the PUR roughness was about 50% that of the wafer. The Bluesil silicone, after curing
in PUR molds and mounting on glass microscope slides, had 75% lower RMS
roughness than the PUR it was cured on. Roughness was low enough that contact
angles on the PDMSe were not significantly affected by the addition of the
nanotopographies (Figure 3-4). The roughness of the Bluesil was also less than that of
the T2 when cured on the same molds (250-2, PUR) (Figure 3-5). The contact angle of
the T2 was increased by the addition of the nanotopography while the contact angle of
the Bluesil was not (Figure 2-7)(Figure 3-4).
51
Assay 1
In the first assay, the settlement density of zoospores was reduced on all
nanotopographies compared to the smooth edges and the entirely smooth slides
(Figure 3-6). Settlement density was reduced by about 50% when comparing the
patterned area to the surrounding smooth area. The completely smooth control slides
had significantly higher settlement density than the smooth areas surrounding the
patterns. Both smooth regions were cured against HMDS coated glass plates, the
contact angles are not significantly different, it is unclear why settlement densities would
be different, more testing is needed. There were also noticeably fewer clumps of spores
on the nanotopographies than on the neighboring smooth areas, it is not known if this is
a result of the pattern or simply of the overall lower number of settled spores. The
reduction of settlement density on the nanotopographies is similar to the reduction seen
during a previous study (Figure 2-8), the first study showed a 74% percent reduction on
the 250-2 topography compared to a 50% reduction in this study. Direct comparisons
should not be made between these studies as different silicones were used (Xiameter
T2 in the previous study and Bluesil RTV 3040 in this one). The new Bluesil does not
replicate or maintain the nanotopography as well as the T2 did, resulting in lower
roughness, this could potentially explain the differences seen in settlement densities on
similar topographies.
Assay 2
A second assay was done with the same topographies and materials to replicate
the results of the first assay. New smooth controls were added to this assay because of
the difference seen in Assay 1 between smooth edge regions and the entirely smooth
slides. The new smooth controls were cut and mounted just like the patterned films. The
52
results from this assay did not match those from Assay 1. Settlement densities were
similar on all topographies, surrounding smooth areas, and most smooth standards
(Figure 3-7). One sample, the smooth PDMSe cured on PUR, showed significantly
lower settlement density than all other samples. This film is cured under the same
conditions and on the same surface as the nanopatterned films, which also showed a
reduction in the first assay, but not the second. This is the first time this sample type has
been tested, it is unclear what cause this reduction or if it is an outlier. Overall
settlement densities were higher on smooth regions in Assay 2 than Assay 1, this
potential difference in spore behavior could account for some of the difference seen
between assays.
Conclusion
The first assay indicated that all the nanotopographies reduced the settlement
density of zoospores by about the same amount. More smooth controls were added to
the second assay to make sure the reduction wasn’t due to another variable introduced
through different curing procedures. The second assay indicated that the
nanotopography had no effect. The results are potentially confounded by the inability of
the silicone to properly replicate the nanotopography, so the effect of the
nanotopography was not truly tested. New surfaces need to be created with different
materials that will replicate topographies better and maintain higher roughness.
Additional testing needs to be done on surfaces of higher roughness to determine if the
nanotopography has any effect on antifouling behavior.
53
Table 3-1. RIE etching parameters for nanopatterned wafers.
RIE Parameters Etch 1 Etch 2 (no particle mask)
Sample RF Power (W) Time (s) RF Power (W) Time (s)
100-1 50 65 50 15
250-1 75 75 75 25
250-2 150 30 150 5
500-1 100 195 n/a n/a
Figure 3-1. SEM micrograph of the fracture surface of silicon wafer with 250-2 topography etched into the top.
54
Figure 3-2. SEM micrographs showing fracture surface of silicon wafers with A) 100-1, B) 250-1, and C) 500-1 topographies etched into the surface.
Figure 3-3. Average RMS roughness data measured with AFM in tapping mode, each
bar is the average of 3 scans (5x5 µm scan size). Error bars represent one standard deviation above and below the mean.
55
Figure 3-4. Average contact angle data of DI water on Bluesil surfaces, each bar represents the average of 5 drops, 5 µL each in volume. Error bars represent one standard deviation above and below the mean.
Figure 3-5. Average RMS roughness of 250-2 nanotopography on PDMSe surfaces
after curing on PUR mold. Error bars represent one standard deviation in each direction.
56
Figure 3-6. The density of spores attached to nanopatterned PDMSe coatings after 45-
minute settlement. Each point is the mean from 90 counts from 3 replicate slides. Bars show 95% confidence limits.
Figure 3-7. The density of spores attached to nanopatterned PDMSe coatings after 45-
minute settlement. Each point is the mean from 90 counts from 3 replicate slides. Bars show 95% confidence limits. Pat is patterned area, Sm is smooth area. Red bars indicate samples cured against PUR, blue bars against glass and the green bar against PFOTS coated wafer.
57
CHAPTER 4 SYLGARD 184 NANOTOPOGRAPHIES FOR ANTIFOULING SILICONE SURFACES
Background
In the previous chapters, PDMSe micro- and nanotopographies were fabricated
for evaluation as marine antifouling coatings. The first study with Xiameter T2 PDMSe
showed potential for the nanoscale topography to decrease the settlement density of
Ulva linza zoospores. Dow Corning discontinued production of the T2 PDMSe,
replacement materials were evaluated for processability and the ability to replicate and
maintain microtopographies. Bluesil RTV 3040 was selected and used for the
fabrication and testing of the nanotopography series. The nanotopography and Ulva
assay results seen with the T2 could not be replicated with the Bluesil. In this study a
new intermediate mold material and a new PDMSe were obtained to create
nanotopographies with higher roughness and to attempt to replicate U. linza assay
results from previous studies.
Objectives
Objective 1: Use new materials and methods to replicate nanotopographies with
higher roughness than Bluesil RTV 3040. In the previous chapter, efforts to replicate
and mount Bluesil topographies resulted in surfaces of low roughness with contact
angle comparable to smooth PDMSe. In this study a new intermediate mold material,
different PDMSe, and different processing procedures will be used to create higher
roughness nanotopographies from the same silicon wafer molds.
Objective 2: Evaluate antifouling potential of PDMSe nanotopographies with U.
linza settlement assay. The settlement assay in this study will use 24-well plates with
sample discs in the bottom of each well instead of mounted on glass microscope slides.
58
This will help minimize contact with the nanotopography during processing to avoid
surface contamination and damage to the topography.
Materials
Wafers, particles, and chemicals were purchased from the same sources as
listed in Chapters 2 and 3. SR415 was obtained from Sartomer (contains Poly(oxy-1,2-
ethanediyl), α-hydro-ω-[(1-oxo-2-propen-1-yl)oxy]-, ether with 2-ethyl-2-
(hydroxymethyl)-1,3-propanediol (3:1) (28961-43-5); < 0.15% 2-Propenoic acid (79-10-
7)). D 1173 curing agent was obtained from BASF, these were used to replace the PUR
as an intermediate mold material. Sylgard 184 PDMSe was purchased from Krayden.
Dow Corning RTV 732 silicone sealant, Corning Costar flat bottom 24-well cell culture
plates (sterile polystyrene), and Fisherbrand aluminum weighing dishes (69 mm wide
and 16 mm deep) were purchased from Fisher Scientific.
Methods
Nanopatterned silicon wafer molds were processed using the same procedures
and parameters described in Chapter 3.
Polymer Replication
Wafers were first treated with PFOTS using the same procedures described in
Chapter 3. Two drops of D 1173 (~40 mg) were added to 4 g of SR 415 in a small glass
vial and mixed thoroughly, any bubbles were then allowed to rise to the surface. A mold
was prepared by attaching a silicon wafer (smooth or with nanotopography) to a smooth
glass plate with double sided tape, with 2mm spacers placed around the edge of the
wafer. A loop of wire the size and shape of the desired SR 415 mold (roughly circular
and 50 mm in diameter) was made and placed gently on the silicon wafer. The wire acts
as a handle for peeling up the mold after polymerization and helps keep the SR415
59
liquid contained before curing. The SR 415 solution was pipetted onto the wafer surface
in and around the wire loop. A transparent quartz plate was then placed on top of the
spacers, compressing and spreading the SR 415. The assembly was then placed in a
UV chamber and illuminated with an ULTRA-VITALUX 300 W 230 V E27 lamp for 25
minutes. After cooling to room temperature, the quartz was carefully separated from the
SR 415 with a scalpel. The solid SR 415 was then peeled off the wafer using the wire
loop handle. The SR 415 molds were placed in the bottom of aluminum weighing dishes
with the nanotopography facing up. Sylgard 184 PDMSe was mixed with a ratio of 10
parts base to 1-part curing agent (by weight), degassed under vacuum, and poured into
the aluminum weighing dish until the whole SR 415 mold is covered by ~2 mm of
PDMSe. The weighing dishes were left in the fume hood to cure at room temperature
for 48 hours. After curing, the PDMSe and SR 415 were removed together from the
aluminum dish, the SR 415 was slowly peeled from the PDMSe.
Bioassay Sample Preparation
A small amount of Dow Corning RTV 732 silicone sealant was spread on the
bottom center of a well in a 24-well plate with a cotton swab (avoiding the edges of the
bottom and the sidewalls). A 14 mm diameter punch was used to punch a round sample
out of the patterned or smooth PDMSe films. Each disc was rinsed with acetone,
ethanol, dried with nitrogen, placed gently into the bottom of a well and pressed gently
around the edges. This procedure was repeated until all wells were full. Care was taken
throughout processing to avoid contact with the central region of the sample discs,
some contact around the edge with tweezers was unavoidable.
60
Ulva linza Settlement Assay
Assays were performed by John Finlay et al. at Newcastle University. Coatings
were equilibrated in 0.22 μm filtered artificial seawater for 24 hours prior to testing.
Zoospores were obtained from mature plants of U. linza by the standard method. A
suspension of zoospores (10 ml; 1x106 spores ml-1) was added to individual wells of the
24-well plate containing the samples. After 45 minutes in darkness at 20 oC, the plates
were emptied and refilled three times in succession to remove unsettled (i.e. swimming)
spores. The spores were fixed using 2.5% glutaraldehyde in seawater. The density of
zoospores attached to the surface was counted by eye using transmitted light
microscopy. Counts were made for 10 fields of view per well (each 0.13 mm2).
Results and Discussion
SR 415 Molds
Silicon wafers were etched using the same procedures as Chapter 3 and the
same topographies were created. Processing of the SR 415 was quicker and more
consistent that the PUR used previously. It can be removed cleanly from a PFOTS
coated wafer and replicated with PDMSe with no further surface treatment. The SR 415
accurately replicated the wafer and exhibited higher RMS roughness than the PUR
embossed on the same wafer mold (Figure 4-1). Figure 4-1A (PUR) has RMS
roughness of 18.3 nm while Figure 4-1B (SR415) has RMS roughness of 26.2 nm. More
important is the quality of replication. The SR 415 mold in Figure 4-1B appears to have
a more consistent top plane, meaning that the SR415 replicated the bottom of the
nanotopography on the wafer accurately than the PUR. The SR 415 molds have higher
RMS roughness than the PUR for the other topographies as well (Figure 4-2), but still
not as high as the wafers. However, the wafer topography consists of raised conical
61
pillars while the PUR and SR415 molds are the inverse of that topography, consisting of
conical pores. This pore structure is more difficult to image accurately than the pillar
topography due to the tip geometry of the AFM tips, especially on the smaller
topographies. If the pore is deeper and higher aspect ratio than the tip, it will not touch
the bottom, but on the wafer images it is clear that the AFM tip touches both the top of
the features and the bottom of the wafer in between them. This could lead to a larger
difference in roughness between the wafer and polymer replicates as measured by
AFM, than actually exists.
Sylgard 184 Topographies
The SG184 PDMSe films tested in this study all have higher RMS roughness
than the Bluesil films tested previously (Figure 4-3). This is also reflected in the contact
angle data (Figure 4-4). While the smaller topographies do not have significantly
different contact angles than the smooth surface, the contact angle on 500-1 is
significantly higher. The contact angle is also higher on the larger SG184 topographies
than on the Bluesil RTV 3040 topographies (Figure 4-4), corresponding to the higher
roughness. While the roughness is higher than the Bluesil topographies, it still does not
fully replicate the roughness of the SR 415 molds or the original wafers.
Ulva linza Settlement Assay
The results of the settlement assay agree with those from the previous chapter
which suggest that there is no difference in settlement between the nanotopographies.
All Sylgard 184 nanotopographies showed the same level of U. linza settlement at
around 400 spores / mm2 (Figure 4-5). In most previous Ulva assays (with the exception
of Assay 2 from Chapter 3) the smooth PDMSe standard showed significantly higher
settlement than any other the patterned surfaces, that was not the case in this assay.
62
The smooth PDMSe performed comparably to the nanotopographies with U. linza
settlement density below 400 spores / mm2. This contrasts with the first assay run in
Chapter 2 in which the smooth sample had settlement density around 1500 spores /
mm2. The polystyrene surface (empty well) showed significantly less settlement than
any of the PDMSe surfaces. Some of these differences could be due to the new assay
type using films in well plates rather than mounted to microscope slides. There was
increased settlement near the edges of the films in the wells, so measurements had to
be taken near the center. A direct comparison between the two assays may not be
possible, more data needs to be collected with this assay type to create a baseline for
comparison.
Conclusion
The combination of new materials, processing methods, and sample
configuration for the Ulva assay resulted in SG184 nanotopographies with higher RMS
roughness than Bluesil RTV 3040 topographies. The roughness of the topography still
decreased with each replication in a new material, though not as much as with PUR and
Bluesil. The contact angle of the larger SG184 topographies was significantly higher
than the Bluesil topographies as well. The Ulva assay data is consistent with the
previous study on Bluesil topographies in that there is no significant difference in
settlement density between any of the nanotopographies, spore counts on topographies
are consistent between studies as well with counts around 400 spores / mm2. There are
however differences in the densities measured on the smooth standards in the different
studies that need to be further investigated.
63
Figure 4-1. AFM height contrast images of A) PUR topography and B) SR 415 topography which were both fabricated on the same 250-1 silicon wafer mold
Figure 4-2. Average roughness of nanotopographies on various materials as measured
by AFM in tapping mode. Averages were taken from 3 images on each surface (5 µm x 5 µm scan size), error bars represent one standard deviation above and below the mean.
64
Figure 4-3. RMS roughness data of final PDMSe sample topographies as measured by
AFM in tapping mode.
Figure 4-4. Average contact angle of 5, 5 µL droplets of DI water on PDMSe surfaces.
Error bars represent 1 standard deviation above and below the mean.
65
Figure 4-5. The density of spores attached to nanopatterned PDMSe coatings after 45-minute settlement. Each point is the mean from 40 counts from 4 replicate wells. Bars show 95% confidence limits.
66
CHAPTER 5
MECHANICAL PROPERTIES OF PDMSe SURFACES
The purpose of this study so far has been to create nanoscale and hierarchical
topographies on PDMSe surfaces to improve that antifouling behavior and extend the
effectiveness to a broader range of organisms. The original silicone resin (Xiameter T2)
that had been previously used and tested for many years by the Brennan Research
Group and others was discontinued by the company and replacement silicones were
found. Unfortunately, the three different silicones tested showed different behavior
during processing, biofouling assays, and in their ability to mold and maintain a
nanotopography. This study will attempt to discover why these silicones behave
differently and which properties are desirable if a new material is chosen to continue this
work in the future.
Background
Silicones have been researched and used for antifouling applications for a long
time due to their low surface energy, low modulus, and ease of processing. Baier’s work
showed that minimal fouling and adhesion of fouling organisms occurred on surfaces
with critical surface tension around 22 mN/m (Baier & DePalma, 1971) (Baier R. , 2006).
Silicones also have surface tension right around that value. The dispersive force of
water’s surface tension is also 22 mN/m so the energy for water to rewet a surface is
minimized when the surface shares this value. A closer examination revealed that this
effect may be more specific to PDMS and its mechanical properties. Other hydrocarbon
and fluorocarbon based polymers tested in the study had more rigid carbon backbones,
but PDMS has a silicon-oxygen backbone with more freedom of rotation and motion
than the carbon-based polymers (Baier & DePalma, 1971) (Brady Jr & Singer, 2000).
67
Another study of surface adhesion used peel tests where a standard 3M Scotch tape
(with acrylic polymer adhesive) was adhered to and peeled off of various surfaces while
the interface was tracked and imaged microscopically. Adhesive fracture energies
measured on the fluorocarbon surfaces were an order of magnitude higher than on the
PDMS surfaces, despite having similar surface free energy (Newby, Chaudhury, &
Brown, 1995) (Zhang Newby & Chaudhury, 1997). A closer analysis of the peel tests
showed relatively large slippage of the adhesive at the PDMS interface. They also
observed less deformation in the bulk of the adhesive while peeling off PDMS than
while peeling off of the fluorocarbon surface showing that most of the shear stress was
concentrated at the PDMS-adhesive interface rather than being distributed through the
bulk. These effects are the result of the unique chemical structure of the siloxane
backbone, which allows for more freedom of motion than a hydrocarbon backbone. This
freedom allows silicone to readily deform and rearrange at a molecular level and as a
bulk material. Unfortunately, these same properties that help prevent fouling and lower
adhesive strength on PDMS and PDMSe surfaces may also hinder the materials ability
to mold and maintain the nanotopographies that the previous chapters have attempted
to test.
Objective
The objective of this study is to identify differences in the mechanical properties
of the silicones tested and determine which properties are desirable in a potential future
replacement material.
Materials
All silicone surfaces used in this study (Xiameter T2, Bluesil RTV 3040, and
Sylgard 184) were retained samples from previous experiments.
68
Methods
Samples were retained from previous experiments, processed using the methods
described in their respective chapters above. AFM analysis was done on a Bruker
Dimension Icon with ScanAsyst in PeakForce Quantitative Nanomechanical Mapping
(PF QNM) mode with ScanAsyst Air tips. In PF QNM mode, a force-distance curve is
collected at each pixel allowing for the imaging of surface topography and the mapping
of multiple other properties simultaneously. All surfaces were imaged using the same
tip, scan parameters and frequency, with a tapping force of 3.5 nN, resulting in
indentations of about 75-100 nm into the surface. The reduced Young’s modulus (E*)
was calculated by the Nanoscope software using the Derjaguin, Muller, Toropov (DMT)
model shown in Equation 5-1, where Ftip is the force on the tip, R is the radius of the
area of the tip touching the sample, d is the tip-sample separation and Fadh is the
adhesion force.
𝐹𝑡𝑖𝑝 =4
3𝐸∗√𝑅𝑑3 + 𝐹𝑎𝑑ℎ (5-1)
Average modulus values and standard deviations represent the average from
262,144 force curves (512 x 512 pixels).
Results and Discussion
The Xiameter T2 PDMSe surface had the highest modulus out of these three
materials averaging 3.4 MPa, compared to 2.17 MPa and 2.67 MPa on Bluesil and
SG184 respectively (Figure 5-1). The T2 was also the silicone that resulted in the
highest RMS roughness when cured on the 250-2 molds (Figure 3-5) (Figure 4-3). The
modulus was also measured on the 250-2 topography as this was the only one used
with all three silicones. The modulus of the T2 was higher on the nanotopography than
69
on the smooth surfaces, while the other two silicones (Bluesil and SG184) had a lower
modulus on the topography than on the smooth surface. The manufacturers reported
shore A hardness values of 42, 38, and 43 for T2, Bluesil, and SG184 respectively;
specific gravity of 1.12, 1.08, and 1.03, and linear shrinkage of < 0.1%. The higher
values of modulus, density and hardness measured on the T2 enhanced the material’s
ability to replicate the nanotopography, as shown by the higher RMS roughness values.
The adhesion data (collected simultaneously with modulus and height data) shows
varying levels of adhesion on the different silicones and topographies from 14-22 nN
(Figure 5-2). The standard deviation of the measured modulus values is much higher on
the topographies than on the smooth surfaces. This seems to be a physical effect
caused by the roughness of the topography because the differing values came from the
tops and bottoms of the features rather than from corresponding locations. All images in
Figure 5-3 were collected simultaneously, the topography is clearly visible in the height
image (Figure 5-3A) and the adhesion image (Figure 5-3B), but not in the modulus
image (Figure 5-3C). According to the height and adhesion images, the lowest adhesion
was measured on the top of the protrusions while the highest adhesion was measured
in between the protrusions. The largest different measured in adhesion between these
locations on the surface was on the Xiameter T2 silicone, which also had the highest
RMS roughness of the 250-2 topographies.
Conclusion
Of the three silicone materials, the Xiameter T2 maintained the highest level of
RMS roughness after curing. This study showed that the T2 also had the highest
surface modulus and density of the silicones tested. If a new silicone is sought to
replace T2 in the creation of nanotopographies it should have a relatively high modulus
70
and density. The potential effect of these mechanical property differences on the fouling
behavior is unclear due to the inconsistency seen in the U. linza settlement assays. Low
modulus has been identified as a potentially desirable property for fouling removal,
which was not tested in these studies. The U. linza assays performed in this were short
term settlement assays to determine the effect of nanotopography on settlement. To
achieve that objective a higher modulus material should be used to accurately
reproduce and test the nanotopographies regardless of the potential effects on fouling
removal.
Figure 5-1. Reduced Young’s modulus as measured by AFM in PF QNM mode. Error
bars represent one standard deviation above and below the mean.
71
Figure 5-2. Adhesive force as measured by AFM in PF QNM mode. Error bars
represent one standard deviation above and below the mean.
Figure 5-3. AFM height (A), adhesion (B), and DMT modulus (C) contrast images.
Images were collected simultaneously in PF QNM mode on the Xiameter T2 250-2 topography.
72
CHAPTER 6 MICROSCALE WETTING BEHAVIOR
Background
While working with microscale silicone surface topographies (such as the
Sharklet AF microtopography and its variations) some observations were made with
implications for the consistency of sample production and testing. As discussed in
previous chapters, the microscale silicone topographies exist in a metastable wetting
state when submerged. Air initially remains trapped in the channels when the sample is
submerged, which means that the wetting state of the surface is sitting in a local
minimum energy state. Energy input is required to overcome the associated energy
barrier and wet the topography (Marmur, 2006). The energy required to partially wet the
topographies used in this study is relatively small as partial wetting has been observed
on some samples after gentle handling and transportation of petri dishes containing
submerged samples. This could cause serious problems experimentally because an
extra variable could be unknowingly added, and the wetting state of each sample could
be unknown and inconsistent at the start of the experiment. This could impact the
results of a biofouling assay because, as discussed in Chapter 2, air that remains
trapped in the topography limits the number of sites available for organisms to settle on
(Decker, et al., 2013). It was observed in the lab that after following the procedures of
collaborating research groups that perform biofouling assays (24-hour soak in ASW),
the wetting state of the samples was inconsistent, and unpredictable without
microscopic evaluation (Figure 6-1). Another reported method to wet topographies
involves a pressurized water jet or squirt bottle, this method was observed to be
inconsistent and unpredictable as well (Figure 6-2). Sonication was tested in this study
73
as an alternative wetting method. It was also observed that some samples had
significant feature flop over before any testing was done, mostly near the edges of the
patterned area (Figure 6-3). This study was performed to understand and prevent the
cause of feature flop over during sample production, and to better understand the
microscale wetting behavior seen on lab scale test samples.
Materials
All materials used in this study and the suppliers are the same as those used in
Chapter 2. Instant Ocean Sea Salt was used to make ASW.
Methods
Wafer patterning and silicone replication were done using the same procedures
described in Chapter 2. Microscope images were taken using a Nikon Eclipse TE2000-
U inverted light microscope. Fifteen different microtopographies were tested, all
variations of the Sharklet AF topography. All topographies were +8.5SK5x5, but 8 of the
patterns maintained the ratio of lengths between neighboring features and therefore the
angles formed by the diamond of the unit cell, these patterns are called the n-series (n2-
n9) based on the number of uniquely sized features in each diamond. The other seven
patterns are called the angle series (a1-a7), they are all n4 patterns with increasing
length difference between neighboring features, changing the angle made by the
diamonds. These patterns were presented and explained by The Brennan Research
Group previously (Decker, 2014). All patterned areas were 2 mm x 2 mm and in line
down the center of each microscope slide so that each slide had one area of each
pattern. Each slide was submerged in a polystyrene petri dish filled with ASW for 24
hours, then imaged (while still submerged) using the inverted microscope to reveal
which areas of the pattern were fully wetted and which still retained air. Some samples
74
were then sonicated in a Fisher Scientific Ultrasonicator in a beaker of ASW and
imaged again. Images were processed using ImageJ to calculate the percent of
patterned area that was wetted.
Results
The first important observation made was the nonuniformity of the patterned
regions. The features near the edges of some patterned areas were all flopped over and
stuck together. The patterned area was also not flat, making it impossible to focus on
the whole region at once with a microscope (Figure 6-3). Through observations after
each stage of processing it became clear that the damage was caused during the
mounting procedure when the patterned silicone film is placed with the patterned side
down on a glass plate, uncured silicone resin is poured on top and the microscope
slides are pressed onto the back with another glass plate. Prior to this step, the patterns
all appeared flat, intact, and with no flop over, so it was not related to the initial curing or
mold release process. The patterned areas of these samples were protruding from the
surrounding surface, so when pressure was applied to the back it was transferred to the
features, causing them to buckle, stick together, and warp the surface. The protruding
patterned areas were also found to influence the wetting behavior of the samples. When
the topographies were submerged, the water was able to immediately flow from the
edge of the pattern, along the long axis of the topography, and fill the channels to where
the first raised features end (Figure 6-4). Through this mechanism, the topography can
become wetted without requiring a Cassie-Baxter to Wenzel transition. While this can
be used to create some interesting wetting patterns (Figure 6-1), it does not accurately
simulate the large patterned area that would be required in a real-world application. This
effect would also not be noticed with contact angle measurements because drops are
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never placed on the edge of a patterned area, they are always placed away from the
edge to better simulate properties of an infinitely large patterned area. A slight change
was made to the pattern writing and etching procedures so that the patterned areas
became recessed in the surrounding surface rather than protruding from it (Figure 6-5).
The spacing between the surrounding border and the ends of the features was kept
consistent with the spacing between features for each pattern to eliminate edge effects
during lab testing.
Another issue which prevented analysis of much of the data was that many of the
images could not be processed in ImageJ after conditioning or after sonication due to
irregular coloration and lighting. Some samples had microscopic or macroscopic
bubbles stuck to the patterned area, obscuring the topography. Since an inverse
microscope was used, any type of contamination or defect on the petri dish, microscope
slide, or PDMSe caused shadows over areas of the topography, making it impossible to
threshold the image and calculate the wetted area (Figure 6-6). Even through
quantification was not possible on many images, it was obvious that sonication was
increasing the wetted area of the topography. Some samples were completely wetted
after 2 min of sonication (Figure 6-7) but others were not (Figure 6-8). Sonication
tended to disperse the air trapped in the channels into smaller bubbles distributed
throughout the topography (Figure 6-8), this made it impossible to see the air
entrapment without microscopic evaluation.
Conclusion
Quantifying the wetted area of most samples under these experimental
parameters was not possible. The qualitative data and observations were instead used
to improve the sample fabrication process. All topographies produced from this point on
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were made to be recessed in the surface rather than protruding, which eliminated most
topographical damage during mounting. Conditioning alone was not sufficient to wet the
topography, nor was squirting with a squirt bottle. Sonication can wet the topography
without damaging the features, but the wetted state cannot be determined by eye, it
must be evaluated microscopically. This should be considered in the future when
conducting testing on wetting or fouling behavior.
Figure 6-1. Inverted optical light microscope images of (A) +8.5SK5x5_n2, (B) +8.5SK5x5_n9, and (C) +8.5SK5x5_n4_a7 PDMSe topographies after 24 hour soak in ASW, surfaces are still submerged in ASW during imaging. The orange areas of the topography are channels that are filled with water, the blue areas are channels that are still filled with air. Patterned areas are 2 mm x 2 mm.
77
Figure 6-2. Inverted light microscope image (unknown magnification) of submerged +3SK2x2_n4 PDMSe topography after squirting surface with squirt bottle while submerged to induce wetting of the topography. Channels are filled with water in the orange areas and air in the blue areas.
Figure 6-3. Inverted light microscope image of +8.5SK5x5_n6 PDMSe topography, the features near the edge look dark because they are laying down and stuck to each other, the central region is out of focus because the patterned area is not flat. The patterned region is 2 mm x 2 mm.
78
Figure 6-4. Inverted light microscope image of submerged +8.5SK5x5_n4_a8 PDMSe topography immediately after submersion in ASW. The orange regions at the edge are channels filled with water, the blue areas still retain air. Patterned area is 2 mm x 2 mm.
Figure 6-5. Schematic representation of (A) “protruding” topography and (B) ”recessed” topography.
79
Figure 6-6. Inverted optical light microscope images of submerged PDMSe
topographies after sonication in ASW. The amount of wetting that has occurred on these surfaces could not be calculated because of bubbles sitting on, below, or within the surface. Patterned areas are 2 mm x 2 mm.
Figure 6-7. Inverted light microscope images of +8.5SK5x5_n9 PDMSe topography after 24 hours of conditioning in ASW (A), and after conditioning and sonication in ASW (B). Patterned area is 2 mm x 2 mm.
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Figure 6-8. Inverted light microscope images of +8.5SK5x5_n8 PDMSe topography after 24 hours of conditioning in ASW (A), and after conditioning and sonication in ASW (B). Patterned area is 2 mm x 2 mm.
81
CHAPTER 7 NANOTEMPLATING WITH BLOCK COPOLYMERS
Background
Block copolymers are a broad class of materials that can be designed and
synthesized for a wide variety of applications. They consist of molecular chains with two
or more segments of different polymers on the same chain. If the chemistry of these
blocks are immiscible in each other they can phase segregate into different
morphologies depending on the molecular weight of each block. For example, if the
molecular weights of the blocks are equal, a lamellar structure may result at equilibrium.
If one block is larger than the other, the minority phase may end up as cylindrical or
spherical domains dispersed in a matrix of the majority block. In a bulk material this
could result in unique properties like the thermoplastic elastomer poly(styrene-block-
butadiene-block-styrene) (SBS). At room temperature polystyrene is below its glass
transition temperature while polybutadiene is above its glass transition temperature.
With a small block of polystyrene at each end of the chain, the resulting bulk polymer at
room temperature has rigid spherical entanglements of polystyrene anchored together
by flexible blocks of polybutadiene. This elastomer can be melted and reformed multiple
times as desired. Block copolymers can also be useful as thin films. With the right
molecular weight ratio, film thickness, and processing methods a morphology can be
obtained where the minority phase consists of an ordered array of cylinders running
through the thickness of the film. If this block is selectively etched away, the result is a
film with an ordered array of nanoscale pores. These films can be used as filtration
membranes, molds for growing nanowires, or in the case of this study as a nanoscale
etching mask. Multiple methods were attempted in this study. Thick films were cast from
82
solutions using methods similar to those used by Sakurai et al., who were able to induce
the coalescence of spherical domains into cylindrical domains oriented perpendicular to
the substrate (Sakurai, et al., 2009). The solvent evaporation from the substrate up
through the film to the free surface creates a chemical potential gradient, which directs
the coalescence of the spherical domains into cylinders during subsequent thermal
annealing. Thin films were produced by spin coating, this is the more traditional method
of obtaining specific domain structures where the thickness of the film relative to the
molecular weight of the polymer chains and blocks can cause perpendicular orientation
of cylindrical domains. Once the domains in the film are oriented, the cylindrical phase
(polybutadiene) can be removed to create an array of nanopores so that the pattern can
be plasma etched into the wafer through the voids in the film. This can be done using
ozone, which reacts with the carbon-carbon double bonds in polybutadiene, breaking it
down into small fragment which are then dispersed with water (Park M. , 1997). After
removal of the polybutadiene, the remaining nanoporous polystyrene film can be used
as an etching mask to transfer the pattern to the underlying silicon wafer.
Materials
Kraton D1403P, D1101, and G1657M were obtained previously as samples from
Kraton and used for solvent casting. P8410-SBdS (SBS) was purchased from Polymer
Source Inc. to be used during spin coating. This material is a triblock copolymer with Mn
of each block of 19,000-b-15,000-b-19,000 and a PDI of 1.07. Toluene, chloroform,
ethanol, and acetone were purchased from Fisher Scientific. One hundred mm and 50
mm diameter, <100> orientation, prime grade single crystal silicon wafers were
purchased from University Wafer. Millipore syringe filters (0.22 µm pore size) were
purchased from Fisher Scientific.
83
Methods
For all casting methods, polymers were dissolved in solvents by slowly adding
the solid polymer pellets to the solvent in a Pyrex bottle while stirring with a magnetic
stir bar on a stir plate.
Solvent Casting Thick Films
Method 1: A 75 mm diameter funnel was inverted and secured over the center of
a smooth or patterned 100 mm silicon wafer. Multiple methods were used to attempt to
secure the wafer including tape and silicone caulk. The polymer and solvent solution
was pipetted into the funnel onto the wafer surface. The whole assembly was left in a
fume hood until all of the solvent was evaporated and a solid film remained on the wafer
surface
Method 2: Two squares of PTFE (75 mm x 75 mm x 12.5 mm) were cut. A 45
mm diameter hole was cut through the center of one PTFE square. A 50 mm silicon
wafer was placed on the solid flat PTFE square, the square with the hole was placed
over the wafer and the PTFE squares were clamped together at the corners using small
C-clamps. A 50 mm diameter funnel was inverted and placed over the hole, the polymer
solution was pipetted onto the wafer. The whole mold was left in a fume hood until the
solvent was evaporated and a solid film remained.
Spin Coating Thin Films
Toluene was filtered through a 0.22 µm syringe filter into a Pyrex bottle. P8410-
SBdS was slowly added to the bottle while stirring. The polymer was allowed to dissolve
completely, then the solution was filtered again and brought to the clean room at the
NRF. A 100 mm silicon wafer was centered on the vacuum chuck of the Headway spin
coater. The wafer was rinsed with acetone, ethanol, and allowed to dry while spinning.
84
One mL of polymer solution was pipetted onto the center of the stationary wafer, the
spin coater speed was then quickly increased to the target max RPM and held for the
desired amount of time. A knob was turned by hand to control speed as this system was
operated manually and was not programmable. The parameters varied during
development of the method were the concentration of P8410-SBdS in the solution and
the max RPM during spin coating.
Etching
An Aqua-6 Multi-Purpose Ozone Generator was purchased from A2Z Ozone, it
can produce 600 mg of ozone per hour. A silicon wafer was coated with SBS using a 3
wt% solution in toluene and spinning at 4000 RPM, it was suspended under DI water in
a glass crystallization dish with the SBS coated side facing down. The ozone generator
connects to a rubber tube with a diffuser stone on the end. The diffuser was secured
below the wafer so that when ozone was produced it bubbled up and across the SBS
coated surface. The generator was run for 20 min/hour, for 24 hours.
Characterization
Thickness measurements were made using a J.A. Woolam ellipsometer. Some
areas of the sample were visibly streaky due to particles present during spin coating,
these areas were avoided during measurement. AFM images were collected using a
Bruker Dimension 3100.
Results
Solvent Casting
The solvent casting method proved to be impractical. When tape was used to
strap the funnel to the wafer, leakage of the solution always occurred. This made it
impossible to cast films with consistent thickness. Caulking between the funnel and
85
wafer prevented leaking, but only because both formed a strong bond with the caulk.
The bond was permanent, preventing further processing of the wafer, like RIE.
Spin Coating
Spin coating with dilute solutions allowed for the casting of films less than 1 µm
thick. The main issue with this technique was consistency. On all wafers that were
coated, particles were present which resulted in streaks in the coating radiating from the
particles towards the edges. These streaks are visible indicators of varying thickness in
the film. Most of the defects occurred near the center of the wafer, which is where any
patterned area would be during processing of a hierarchical structure. To prevent
particle contamination the rest of the process would need to be moved into the clean
room, but this would still not guarantee a uniform coating. The vacuum chucks on this
spin coater also did not sit perfectly level on the spindle, causing the wafer to wobble
while spinning. Areas on the wafer which appeared streak free and uniform were
measured using ellipsometry which showed that thickness in these areas could still vary
by tens of nanometers, which could be greater than 10% of the average film thickness.
To attain the desired block copolymer morphology, specific and consistent film
thickness is required. The variation measured using his method makes that impossible.
All of these measurements were taken on smooth wafers, spin coating on wafers with
microtopographies etched into them would add even more challenges to the process.
Etching
Based on AFM imaging, the ozone treatment appeared to successfully remove
the polybutadiene blocks. AFM images taken on the film surface before ozone treatment
showed a relatively smooth surface with no observable nanoscale phase segregation
(Figure 7-1). Images after treatment revealed a phase segregated morphology was
86
present, but it was either under a surface layer of polybutadiene, or was not observable
in height contrast images (Figure 7-2). The remaining film on the wafer was a
nanoporous polystyrene film. While this was the desired result, the desired morphology
was not obtained. There was no ordered packing structure to the pores, nor were they
all round. Also, while this was close to the desired result, the AFM images shown are
not representative of the structure across the entire wafer. As mention previously there
were large visible defects in the spin coated films, the measurements and images
discussed here represent only the best areas of the coating.
Conclusion
Block copolymers can be used to create nanoporous films, and while solvent
casting and spin coating are both possible routes to achieve this goal, neither method
was practical for this study. Using the facilities available, and due to the relatively large
area of coating required for future testing, consistent coating thickness could not be
obtained. With all the difficulties encountered working with smooth wafers, it quickly
became obvious that a new nanopatterning method would be needed to create
hierarchical structures on micropatterned wafer.
87
Figure 7-1. AFM height contrast images of SBS surface with no apparent organized nanoscale phase segregation
Figure 7-2. AFM height contrast images of SBS surface after ozone removal of polybutadiene. Areas previously filled with polybutadiene now appear as dark holes in the remaining polystyrene film.
88
CHAPTER 8 SUMMARY AND FUTURE WORK
Observations of the microscale wetting behavior of lab scale samples led to the
successful revision of fabrication procedures to prevent damage during fabrication and
edge effects during testing. Nanoscale and hierarchical topographies have been
successfully fabricated on silicon wafers through multiple coating and etching
processes. These topographies were replicated in PDMSe with varying degrees of
success depending on the materials and methods used. The nanotopography molded
into the first PDMSe that was tested (Xiameter T2) showed promise in reducing the
settlement of U. linza zoospores. Unfortunately, these samples could not be retested
due to the discontinuation of the Xiameter T2 by the manufacturer. The Bluesil RTV
3040 which was obtained as a replacement silicone could not replicate and maintain the
nanotopography as well as the Xiameter T2, so the bioassay results were not really a
test of the nanotopographies as was desired. The new materials, replication and
mounting process used in Chapter 4 resulted in samples of higher roughness, but still
not as high as seen on the original wafer molds or the Xiameter T2 samples, and there
was still no difference seen in the settlement of the U. linza zoospores on the
nanotopographies. Other parameters varied between the multiple silicone materials
tested including the modulus and density, this not only affects the ability of the material
to replicate the nanotopography, but could confound results on the different smooth
surfaces tested as well. It would not be wise to make direct comparisons between the
different assays due to these confounding variables, especially with the high variation
typically associated with biological data even when sample properties are consistent.
89
Should this project be continued in the future, a new, more suitable PDMSe
should be found first. This material should have a higher modulus and density than the
silicones tested so far in this study. Different nanotopography morphologies and aspect
ratios may also help the PDMSe maintain higher roughness. PDMSe topographies
should not be mounted on glass microscope slides before testing. Future bioassays
should all be conducted using the well plate method if possible. This will help keep
topographies intact and sample surfaces as pristine as possible by minimizing contact
with other surfaces.
90
LIST OF REFERENCES
Alzieu, C., Sanjuan, J., Deltreil, J., & Borel, M. (1986, 11 1). Tin contamination in Arcachon Bay: Effects on oyster shell anomalies. Marine Pollution Bulletin, 17(11), 494-498.
Baier, R. (1981). Early events of micro-biofouling of all heat transfer equipment. In R.
Baier, Fouling of heat transfer equipment (pp. 293-304). Baier, R. (2006). Surface behaviour of biomaterials: The theta surface for
biocompatibility. Journal of Materials Science: Materials in Medicine, 17(11), 1057-1062.
Baier, R., & DePalma, V. (1971). Management of occlusive arterial disease. Yearbook
Medical, Chicago, 147. Baier, R., & DePalma, V. (1971). The relation of the internal surface of grafts to
thrombosis. Management of Arterial Occlusive Disease, 18, 1-47. Bandara, C., Singh, S., Afara, I., Wolff, A., Tesfamichael, T., Ostrikov, K., & Oloyede, A.
(2017, 3 16). Bactericidal Effects of Natural Nanotopography of Dragonfly Wing on <i>Escherichia coli</i>. ACS Applied Materials & Interfaces, 9(8), 6746-6760.
Bers, A., & Wahl, M. (2004, 2). The Influence of Natural Surface Microtopographies on
Fouling. Biofouling, 20(1), 43-51. Borkow, G., & Gabbay, J. (2009, 9 1). Copper, An Ancient Remedy Returning to Fight
Microbial, Fungal and Viral Infections. Current Chemical Biology, 3(3), 272-278. Brady Jr, R., & Singer, I. (2000). Mechanical factors favoring release from fouling
release coatings. Biofouling: The Journal of Bioadhesion and Biofilm Research Biofouling, 15(15), 1-3.
Callow, M. (1990). Ship fouling. Problems and solutions (Vol. 5). Callow, M., Jennings, A., Brennan, A., Seegert, C., Gibson, A., Wilson, L., . . . Callow, J.
(2002). Microtopographic cues for settlement of zoospores of the green fouling alga <i>Enteromorpha</i>. Biofouling, 18(3), 229-236.
Cao, X., Pettitt, M., Wode, F., Sancet, M., Fu, J., Jian, J., . . . Grunze, M. (2010).
Interaction of zoospores of the green alga ulva with bioinspired micro- And nanostructured surfaces prepared by polyelectrolyte layer-by-layer self-assembly. Advanced Functional Materials, 20(12), 1984-1993.
Carman, M., Estes, T., Feinberg, A., Schumacher, J., Wilkerson, W., Wilson, L., . . .
Brennan, A. (2006). Engineered antifouling microtopographies - correlating wettability with cell attachment. Biofouling, 22(1), 11-21.
91
Carr, L., Xue, H., & Jiang, S. (2011). Functionalizable and nonfouling zwitterionic
carboxybetaine hydrogels with a carboxybetaine dimethacrylate crosslinker. Biomaterials, 32(4), 961-968.
Cassie, A., & Baxter, S. (1944, 1 1). Wettability of porous surfaces. Transactions of the
Faraday Society, 40(0), 546. Dafforn, K., Lewis, J., & Johnston, E. (2011, 3 1). Antifouling strategies: History and
regulation, ecological impacts and mitigation. Marine Pollution Bulletin, 62(3), 453-465.
Davidson, I., Brown, C., Sytsma, M., & Ruiz, G. (2009). The role of containerships as
transfer mechanisms of marine biofouling species. Biofouling, 25(7), 645-655. Decker, J. (2014). A Predictive Model for the Antifouling Efficacy of Engineered
Microtopographies. Doctoral DIssertation. Decker, J., Kirschner, C., Long, C., Finlay, J., Callow, M., Callow, J., & Brennan, A.
(2013). Engineered antifouling microtopographies: An energetic model that predicts cell attachment. Langmuir, 29(42).
Ecol Prog Ser, M., & Wahl, M. (1989). MARINE ECOLOGY PROGRESS SERIES
Marine epibiosis. I. Fouling and antifouling: some basic aspects. 58, 175-189. Ekblad, T., Bergström, G., Ederth, T., Conlan, S., Mutton, R., Clare, A., . . . Liedberg, B.
(2008). Poly(ethylene glycol)-containing hydrogel surfaces for antifouling applications in marine and freshwater environments. Biomacromolecules, 9(10), 2775-2783.
Greco, G., Lanero, T., Torrassa, S., Young, R., Vassalli, M., Cavaliere, A., . . .
Davenport, J. (2013). Microtopography of the eye surface of the crab Carcinus maenas: an atomic force microscope study suggesting a possible antifouling potential. Journal of the Royal Society Interface, 10(84).
Hasibeck, E., & Bohlander, G. (1992). Microbial Biofilm Effects on Drag -Lab and Field.
Proceedings of the Ship Production Symposium (pp. 2-4). New Orleans: THE SOCIETY OF NAVAL ARCHITECTS AND MARINE ENGINEERS.
Hoipkemeier-Wilson, L., Schumacher, J., Carman, M., Gibson, A., Feinberg, A., Callow,
M., . . . Brennan, A. (2004, 2). Antifouling Potential of Lubricious, Micro-engineered, PDMS Elastomers against Zoospores of the Green Fouling Alga <i>Ulva (Enteromorpha)</i>. Biofouling, 20(1), 53-63.
92
Holland, R., Dugdale, T., Wetherbee, R., Brennan, A., Finlay, J., Callow, J., & Callow, M. (2004, 9). Adhesion and motility of fouling diatoms on a silicone elastomer. Biofouling, 20(6), 323-329.
Hu, Z., Finlay, J., Chen, L., Betts, D., Hillmyer, M., Callow, M., . . . Desimone, J. (2009).
Photochemically cross-linked perfluoropolyether-based elastomers: Synthesis, physical characterization, and biofouling evaluation. Macromolecules, 42(18), 6999-7007.
Hui, C.-Y., & Jagota, A. (2014, 5 7). Deformation near a liquid contact line on an elastic
substrate. Proceedings of the Royal Society A: Mathematical, Physical and Engineering Sciences, 470(2167), 20140085-20140085.
Jeon, S., Lee, J., Andrade, J., & De Gennes, P. (1991). Protein-surface interactions in
the presence of polyethylene oxide. I. Simplified theory. Journal of Colloid And Interface Science, 142(1), 149-158.
Jerison, E., Xu, Y., Wilen, L., & Dufresne, E. (2011). The Deformation of an Elastic
Substrate by a Three-Phase Contact Line. Phys. Rev. Lett, 106(18), 186103. Jiang, S., & Cao, Z. (2010). Ultralow-fouling, functionalizable, and hydrolyzable
zwitterionic materials and their derivatives for biological applications. Advanced Materials, 22(9), 920-932.
Kelleher, S., Habimana, O., Lawler, J., O'reilly, B., Daniels, S., Casey, E., & Cowley, A.
(2016). Cicada Wing Surface Topography: An Investigation into the Bactericidal Properties of Nanostructural Features. ACS Applied Materials and Interfaces, 8(24).
Kempf, G. (1937). On the effect of roughness on the resistance of ships. Trans INA, 79,
109-119. Kesel, A., & Liedert, R. (2007). Learning from Nature: Non-Toxic Biofouling Control by
Shark Skin Effect (Vol. 146). Koch, K., Bhushan, B., & Barthlott, W. (2009, 2). Multifunctional surface structures of
plants: An inspiration for biomimetics. Progress in Materials Science, 54(2), 137-178.
Krishnan, S., Ayothi, R., Hexemer, A., Finlay, J., Sohn, K., Perry, R., . . . Fischer, D.
(2006). Anti-biofouling properties of comblike block copolymers with amphiphilic side chains. Langmuir, 22(11), 5075-5086.
Krishnan, S., Wang, N., Ober, C., Finlay, J., Callow, M., Callow, J., . . . Fischer, D.
(2006). Comparison of the fouling release properties of hydrophobic fluorinated
93
and hydrophilic PEGylated block copolymer surfaces: Attachment strength of the diatom Navicula and the green alga Ulva. Biomacromolecules, 7(5), 1449-1462.
Leh, A., N’guessan, H., Fan, J., Bahadur, P., Tadmor, R., & Zhao, Y. (2012, 4 3). On the
Role of the Three-Phase Contact Line in Surface Deformation. Langmuir, 28(13), 5795-5801.
Lewis, J. (1998). Marine Biofonling and its Prevention on Underwater Surfaces.
Materials Forum, 22, pp. 41-6I. Marabotti, I., Morelli, A., Orsini, L., Martinelli, E., Galli, G., Chiellini, E., . . . Jenko, M.
(2009). Fluorinated/siloxane copolymer blends for fouling release: chemical characterisation and biological evaluation with algae and barnacles. Biofouling, 25(6), 481-493.
Marmur, A. (2006). Super-hydrophobicity fundamentals : implications to biofouling
prevention. Biofouling, 22(September 2013), 37-41. Martinelli, E., Suffredini, M., Galli, G., Glisenti, A., Pettitt, M., Callow, M., . . . Lyall, G.
(2011). Amphiphilic block copolymer/poly(dimethylsiloxane) (PDMS) blends and nanocomposites for improved fouling-release. Biofouling, 27(5), 529-541.
Newby, B.-m., Chaudhury, M., & Brown, H. (1995, 9 8). Macroscopic Evidence of the
Effect of Interfacial Slippage on Adhesion. Science, 269(5229), 1407. Omae, I. (2003, 2 1). Organotin antifouling paints and their alternatives. Applied
Organometallic Chemistry, 17(2), 81-105. Ostuni, E., Chapman, R., Holmlin, R., Takayama, S., & Whitesides, G. (2001). A survey
of structure-property relationships of surfaces that resist the adsorption of protein. Langmuir, 17(18), 5605-5620.
Otani, M., Oumi, T., Uwai, S., Hanyuda, T., Prabowo, R., Yamaguchi, T., & Kawai, &.
(2007). Occurrence and diversity of barnacles on international ships visiting Osaka Bay, Japan, and the risk of their introduction. Biofouling, 23(4), 277-286.
Pangam, P., Giriyan, A., & Hawaldar, K. (2009, 2 15). Implications of the ban on
organotins for protection of global coastal and marine ecology. Journal of Environmental Management, 90, S96-S108.
Park, D., Weinman, C., Finlay, J., Fletcher, B., Paik, M., Sundaram, H., . . . Ober, C.
(2010). Amphiphilic surface active triblock copolymers with mixed hydrophobic and hydrophilic side chains for tuned marine fouling-release properties. Langmuir, 26(12), 9772-9781.
94
Park, M. (1997). Block Copolymer Lithography: Periodic Arrays of 10^11 Holes in 1 Square Centimeter. Science, 276(5317), 1401-1404.
Piola, R., Dafforn, K., & Johnston, E. (2009). The influence of antifouling practices on
marine invasions. Biofouling The Journal of Bioadhesion and Biofilm ResearchOnline) Journal, ISSN homepage, 892-7014.
Pogodin, S., Hasan, J., Baulin, V., Webb, H., Truong, V., Nguyen, H., . . . Ivanova, E.
(2013). Biophysical Model of Bacterial Cell Interactions with Nanopatterned Cicada Wing Surfaces. Biophysj, 104, 835-840.
Sakurai, S., Bando, H., Yoshida, H., Fukuoka, R., Mouri, M., Yamamoto, K., &
Okamoto, S. (2009). Spontaneous perpendicular orientation of cylindrical microdomains in a block copolymer thick film. Macromolecules, 42(6), 2115-2121.
Scardino, A., Guenther, J., & de Nys, R. (2008, 1). Attachment point theory revisited:
the fouling response to a microtextured matrix. Biofouling, 24(1), 45-53. Scardino, A., Hudleston, D., Peng, Z., Paul, N., & de Nys, R. (2009, 1). Biomimetic
characterisation of key surface parameters for the development of fouling resistant materials. Biofouling, 25(1), 83-93.
Schultz, M., Bendick, J., Holm, E., & Hertel, W. (2011). Economic impact of biofouling
on a naval surface ship. Biofouling, 27(1), 87-98. Schumacher, J., Aldred, N., Callow, M., Finlay, J., Callow, J., Clare, A., & Brennan, A.
(2007). Species-specific engineered antifouling topographies: correlations between the settlement of algal zoospores and barnacle cyprids. Biofouling, 23(5), 307-317.
Schumacher, J., Carman, M., Estes, T., Feinberg, A., Wilson, L., Callow, M., . . .
Brennan, A. (2007). Engineered antifouling microtopographies - effect of feature size, geometry, and roughness on settlement of zoospores of the green alga Ulva. Biofouling, 23(1), 55-62.
Seo, K., Kim, M., & Kim, D. (2015). Re-derivation of Young’s Equation, Wenzel
Equation, and Cassie-Baxter Equation Based on Energy Minimization. In K. Seo, M. Kim, D. Kim, & M. Aliofkhazraei (Ed.), Surface Energy (p. Ch. 01). Rijeka: InTech.
Style, R., Che, Y., Wettlaufer, J., Wilen, L., & Dufresne, E. (2013). Universal
deformation of soft substrates near a contact line and the direct measurement of solid surface stresses. Phys. Rev. Lett, 110(6), 066103.
Townsin, R. (2003). The Ship Hull Fouling Penalty. Biofouling, 19(sup1), 9-15.
95
Turner, A. (2010, 2 1). Marine pollution from antifouling paint particles. Marine Pollution
Bulletin, 60(2), 159-171. Weinman, C., Gunari, N., Krishnan, S., Dong, R., Paik, M., Sohn, K., . . . Ober, C.
(2010). Protein adsorption resistance of anti-biofouling block copolymers containing amphiphilic side chains. Soft Matter, 6(14), 3237.
Wenzel, R. (1936). Resistance of solid surfaces to wetting by water. Journal of Industrial
and Engineering Chemistry (Washington, D. C.), 28, 988-994. WHOI. (1952). Marine fouling and its prevention ; prepared for Bureau of Ships, Navy
Dept. Woods Hole, MA: United States Naval Institute. Yang, H., Dou, X., Fang, Y., & Jiang, P. (2013, 9 1). Self-assembled biomimetic
superhydrophobic hierarchical arrays. Journal of Colloid and Interface Science, 405, 51-57.
Yang, K., Jung, K., Ko, E., Kim, J., Park, K., Kim, J., & Cho, S.-W. (2013).
Nanotopographical Manipulation of Focal Adhesion Formation for Enhanced Differentiation of Human Neural Stem Cells. ACS Applied Materials & Interfaces, 5(21), 10529-10540.
Zhang Newby, B.-m., & Chaudhury, M. (1997, 3 1). Effect of Interfacial Slippage on
Viscoelastic Adhesion. Langmuir, 13(6), 1805-1809. Zhang, Z., Finlay, J., Wang, L., Gao, Y., Callow, J., Callow, M., & Jiang, S. (2009).
Polysulfobetaine-grafted surfaces as environmentally benign ultralow fouling marine coatings. Langmuir, 25(23), 13516-13521.
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BIOGRAPHICAL SKETCH
Clayton Walker Argenbright was born in Longwood, Florida in 1990 to Janet and
Michael Argenbright. He grew up mainly in the areas surrounding the Wekiva Springs
State Park, exploring and enjoying Florida’s natural areas. He graduated from Lake
Brantley High School in 2008 and moved to Gainesville to attend the University of
Florida, intent on studying engineering.
As an undergraduate he developed an interest in materials science and began
his research under Dr. Scott Perry, learning how to build and operate XPS systems. He
remained at the University of Florida to continue to study with Dr. Anthony Brennan in
the field of antifouling surface topographies. When not working in a lab, Clayton could
be found outside; fishing, floating down a river, or on a disc golf course.