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

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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.

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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

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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

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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.

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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.

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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.

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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.

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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

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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.

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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

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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.