thesis-farida yasmin-final

Dependency of Cell Mechanics on Substrate Topography

Dissertation submitted in partial fulfilment of the requirements for the degree of “Master of Science” (MSc) of the Program of Advanced Materials Ulm University

Submitted by

Farida Yasmin 849697 [email protected] Dhaka June 2016

mailto:[email protected]

Head of the Department

Prof Dr. Kay E. Gottschalk

First Supervisor:

Prof Dr. Kay E. Gottschalk

University Ulm

Second Supervisor:

Prof Dr. Rolf Brenner

Universitätsklinikum Ulm

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Contents

Abbreviations …………………………………………………………………………………………………………………………..10

Abstract ……………………………………………………………………………………………………………………………………11

Chapter 1. Introduction ............................................................................................................ 12

Materials Science And Engineering ................................................................................ 12

1.1.1 Nanomaterials ........................................................................................................ 12

1.1.2 Biomaterials ............................................................................................................ 13

Importance of Studying Cell Mechanics In Biomaterials ................................................ 14

Chapter 2. Background Information......................................................................................... 15

An Introduction To Cell ................................................................................................... 15

Cell –Surface interaction ................................................................................................ 17

Theory of Elasticity (Hertz Contact Model) .................................................................... 18

Chapter 3. Experimental........................................................................................................... 20

Cleanroom Technique .................................................................................................... 20

Spin Coating: Deposition By Spinning ............................................................................. 21

Dip Coating: Preparation of Nanoparticles ..................................................................... 22

H2 Plasma ........................................................................................................................ 24

Photochemical Deposition ............................................................................................. 25

Plasma Etching: Reactive Ion Etching (RIE) .................................................................... 26

3.6.1 Effects of Oxygen addition ..................................................................................... 28

3.6.2 Effects of Hydrogen addition .................................................................................. 28

3.6.3 Effects of CHF3 and Noble gas addition .................................................................. 28

Electron Microscope ....................................................................................................... 29

3.7.1 Scanning Electron Microscopy (SEM) ..................................................................... 29

3.7.2 Atomic Force Microscopy (AFM) ............................................................................ 30

Cell Culture Preparation ................................................................................................. 34

Chapter 4. Results and Discussion ............................................................................................ 35

Fabrication of Nano-Pillars ............................................................................................. 35

4.1.1 Micellar technique by using block copolymer ........................................................ 35

4.1.2 Photochemical growth of Gold (Au) particle .......................................................... 37

4.1.3 Reactive Ion Etching (RIE) ....................................................................................... 38

Cell Mechanics on Nanostructure Topography .............................................................. 41

4.2.1 Indentation depth ................................................................................................... 41

4.2.2 Measurements........................................................................................................ 42

Cell-Surface interaction .................................................................................................. 53

Chapter 5. Conclusion .............................................................................................................. 57

Chapter 6. References .............................................................................................................. 58

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List of Tables

Table 3-1 Preparation of micellar solution ..................................................................................... 23

Table 3-2 Description of different types of filter ............................................................................ 24

Table 3-3 Preparation of the gold solution for photochemical growth ......................................... 26

Table 3-4: Settings of AFM ............................................................................................................. 33

Table 4-1: Experimental data of particle diameter, etching time and average height .................. 40

Table 4-2: Elastic modulus with different Samples and Spring Constant ....................................... 53

List of Figures

Figure 1-1: Correlation of nano and biotechnology [8] .................................................................. 13

Figure 2-1: Schematic diagram of a Eukaryotic cell [18] ................................................................ 15

Figure 2-2: Schematic diagram of Fibroblast in ECM [22] .............................................................. 16

Figure 2-3: Schematic diagram of components of cytoskeleton, a) Microtubules b) Intermediate

filament c) Actin filament [25] ....................................................................................................... 17

Figure 2-4: Movement of cell by crawling over the surface [28] ................................................... 17

Figure 2-5: Schematic representation of a cell possessing different types of forces [30] ............. 18

Figure 2-6: Schematic diagram of Hertz contact model with a spherical tip (radius R2), loading

force F, Cell radius R1, a is contact radius and δ is indentation depth [34] .................................... 19

Figure 3-1: Classification of cleanroom [39] ................................................................................... 20

Figure 3-2: schematic diagram of different steps of spin coating: a) dispensation of photoresist,

b) acceleration, c) spreading of the liquid, d) evaporation [40] ..................................................... 21

Figure 3-3: Schematic diagram of stages of dip-coating process [46] ............................................ 22

Figure 3-4: Preparation of Au micellar solution [50] ...................................................................... 23

Figure 3-5: Structure of PS-b-P2VP blocked copolymer, modified from [51] ................................ 23

Figure 3-6 Photochemical growth of Au particle [54] .................................................................... 25

Figure 3-7: Schematic diagram of optical system of mask aligner Karl SUSS MJB 3 Mask UV 400

[58] ................................................................................................................................................. 26

Figure 3-8: Schematic representation of the process of etching adopted from [61] .................... 27

Figure 3-9: Schematic diagram of Scanning Electron Microscope (SEM) [66] ............................... 29

Figure 3-10: Emission of various electrons and electromagnetic waves from the specimen [65] 30

Figure 3-11: Schematic diagram of AFM [69] ................................................................................. 31

Figure 3-12: Schematic diagram of scanning system in AFM, redrawn from [70] ......................... 31

Figure 3-13: Force spectroscopy mode in AFM [71] ...................................................................... 32

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Figure 3-14: Calibration of cantilever spring constant, blue line is for cantilever resonance peak

(measured) and red is the Lorentz fit [71] ..................................................................................... 33

Figure 4-1: Plot of average interparticle distance over withdrawal velocity from experimental

data, from figure it is seen that for 130nm interparticle distance the withdrawal speed is

2.8mm/min ..................................................................................................................................... 36

Figure 4-2: Interparticle distance changes depending on the concentration of the solution [58] 36

Figure 4-3: HRSEM image of Au nanoparticle after H2 Plasma. A) Grey scale picture and B) after

adjusting threshold, inset hexagonal arrangement (bandpass filter). Scale is 1µm, 30kV. ........... 36

Figure 4-4: Plot of Au NPs diameter as a function of exposure time from experimental data. ..... 37

Figure 4-5: HRSEM images (at 30kV) of different diameter(average) of Au NPs, a) 9nm, b) 17nm,

c) 30nm, scale is 200nm. ................................................................................................................ 37

Figure 4-6: a) grey scale image using HRSEM scale 200nm at 30kV; b) after adjusting threshold;

and c) marking for area measurement through imageJ software ................................................. 38

Figure 4-7: Schematic diagram of a) formation of SiF2 and b) formation of SiF4 [79] ................... 38

Figure 4-8: Etching rate changes depending on a) DC bias and b) size of the mask [77] ............... 39

Figure 4-9: Au NPs in 200nm scale, tilted by 30 degree a) average height 75nm and diameter

28nm with Au particle diameter 17nm, b) height 75nm and diameter 49nm with Au particle

diameter 30nm, and c) height 106nm and diameter 49nm with Au particle diameter 30nm ...... 39

Figure 4-10: AFM pictures (topography) of Samples 2 (130-28-75) ............................................... 40

Figure 4-11: Height of pillars build by Gwydion, profile 1 corresponds to red line and profile 2

corresponds to yellow line from Figure 4-10 right part. ................................................................ 41

Figure 4-12: Schematic diagram of indentation test (top) and force-indentation curve (bottom)

[14] ................................................................................................................................................. 41

Figure 4-13: Sample 1: 130-49-106, Hertzfit indentation depth 100nm, figure A shows the

topography of the cell, C is force-distance curve, B is ‘Young’s modulus error’ vs ‘indentation

depth’ and D is ‘Young’s modulus ‘vs ‘Indentation depth’ curve ................................................... 42

Figure 4-14: Histogram of Young’s modulus by using Hertz model of 3T3 fibroblast of Sample 1 at

indentation depth 100nm. X-axis is in logarithm, Y-axis is linear scale (calibrated spring constant

0.275 N/m), different colours produced by the addition of one cell with other and this results

come from summation of 10 cells data. ......................................................................................... 43

Figure 4-15: Histogram of Sample 2 at indentation depth 100nm 30 cells measurement,

(calibrated spring constant-0.125N/m), different colours arise by adding one cell with other .... 44

Figure 4-16: Histogram of sample 3a at indentation depth 100nm, with spring constant 0.084

N/m, different colours arise from summation of all cells (6cells measurement). ......................... 45

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Figure 4-17: Histogram of Sample 3b at indentation depth 100nm (5cells measurement) with

calibrated spring constant 0.275 N/m, different colours arise from summation of all cells. ........ 45

Figure 4-18: Histogram of Sample 1 (top, 10 cells) and Sample 3b (bottom, 5 cells), frequency

scale is different due to the different number of cells measurements, calibrated spring constant

0.275 N/m. Different colours produced by the addition of one cell with other. ........................... 46

Figure 4-19: Boxplot for comparing the median of different samples at indentation depth-100nm

........................................................................................................................................................ 47

Figure 4-20: All samples: Sample 1: 130-49-106, Sample 2: 130-28-75, Sample 3a: 130-49-75 and

Sample 3b:130-49-75 ..................................................................................................................... 48

Figure 4-21: Boxplot of all samples of Young’s modulus over selected height: 0.4-0.6, 0.9-1.1, 1.4-

1.6 and 1.9-2.1 µm ......................................................................................................................... 49

Figure 4-22: Young’s modulus VS Height plot of 3 samples: Sample1: 130-49-106, Sample2: 130-

28-75, and Sample3b: 130-49-75 ................................................................................................... 49

Figure 4-23: Boxplot of 3 samples of Young’s modulus over selected height, the selected heights

are: 0.4-0.6, 0.9-1.1, 1.4-1.6 and 1.9-2.1 µm ................................................................................. 50

Figure 4-24: 3D plot of Young’s modulus over height of 3 samples, colour scale bar shows how

frequent the combination of young’s modulus and height was measured, left figures are for the

cell and right figures are for the substrate. .................................................................................... 52

Figure 4-25: AFM picture (topography) of Sample 1 after background corrections, scale 5µm,

colour scale bar shows the height measured ................................................................................. 54

Figure 4-26:HRSEM pictures at 5kV, Cells spread over the surface of all types of samples; a)

Elongated triangular shape with formation of microvili, scale 20µm, b) without microvili, scale

30µm c) the formation of lamellipodia and filopodia (yellow arrows), scale 5µm ........................ 55

Figure 4-27: HRSEM picture taken at 5kV, 30 degree tilted, Cell moves attaching the top of the

pillars in all samples (black arrows) a) Sample 2 scale 100nm, b) Sample 1, scale 1µm and c)

Sample 3b 1µm. .............................................................................................................................. 56

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‘Everybody is a genius. But, if you judge a fish by its ability to climb a tree, it will spend its

whole life believing that it is stupid.’

Albert Einstein

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This work is dedicated to my beloved husband who has always inspired me,

supported me and encouraged me. Also to my parents and my little sister

who have always prayed for me and my brother for guiding me.

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DECLARATION

I hereby declare that I wrote the present dissertation with the topic: Dependency of Cell

Mechanics on Substrate Topography independently and used no other aids than those cited. In

each individual case, I have clearly identified the source of the passages that are taken word for

word or paraphrased from other works. I also hereby declare that I have carried out my

scientific work according to the principles of good scientific practice in accordance with the

current “Satzung der Universität Ulm zur Sicherung guter wissenschaftlicher Praxis“[Rules of the

University of Ulm for Assuring Good Scientific Practice].

…………………………..

Farida Yasmin

Ulm, 10.06.2016

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ACKNOWLEDGEMENT

I’m obliged to my first supervisor Prof. Dr. Kay E. Gottschalk to give me this opportunity to work

in a wide field of Biomaterials & Biophysics, also I show my gratitude to Prof. Dr. Rolf Brenner to

be my second supervisor. My deepest respect to my Ex-supervisor Dr. Alfred Plettl, I’m grateful

to Dr. Axel Seidenstüker to train me well in Cleanroom Technique, Scanning Electron Microscopy

and for being nice to me. I show courtesy to Fabian Endele, Tanja and Anja for helping me during

my work. I’m thankful to Patrick Paul to introduce me to Atomic Force Microscopy. I mostly

grateful to Nicole Sieber for helping me a lot with measurements. I want to thank Ulla Nolte to

do the cell culture for me. I want to thank my husband Dr. Mohammad Abbas Uddin, my friend

Sean Harvey and Dr. Nabiul Hassan, and Patrick Paul for being my proof reader.

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ABBREVIATIONS

AFM Atomic Force Microscopy

Au NPs Gold nanoparticles

ECM Extracellular matrix

HRSEM High Resolution Scanning Electron Microscopy

PMMA Poly methyl methacrylate

P2VP Poly(2-vinylpyridine)

PS-b-P2VP

BCMT

Polystyrene-block-Poly-2-vinylpyridin

Block Copolymer Micellar Technique

RIE

Silicon

F

Ar

He

H

HF

CF4

CHF3

E

3T3

Sccm

Reactive Ion Etching

Si

Fluorine

Argon

Helium

Hydrogen

Hydrogenflouride

Tetrafluoromethane

Trifluoromethane

Elastic constant

3-day transfer, inoculum 3 x 105 cells

Standard Cubic centimetres

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ABSTRACT

Mechanical property of materials behave differently when they are subjected to another

materials with different topography, particularly for biological cells. Cytoskeleton and nucleus,

components of biological cells, varies in their mechanical properties such as cell stiffness and

rigidity when another material is applied. Also, depending on the topography cell stiffness

changes. In this regard, the nature of interaction between soft materials like cell to hard

materials like Silicon is worth exploratory. Silicon has been used as a biomaterials for long

however, very few researches were carried out to measure the mechanical properties of

fibroblast cells on nanostructure of Silicon. Therefore, this study investigated the mechanical

property of 3T3 fibroblast (mouse embryonic dermal cell) on Silicon nanostructure surface and

corresponding interaction between them. In this work Silicon nano-pillars were fabricated from

n-type Silicon wafer with different heights and diameters.

Characterization of the fabricated nano-pillar surface was carried out with HRSEM (High

Resolution Scanning Electron Microscopy), and the elasticity of cell on substrate was measured

by Atomic Force Microscopy (AFM). It was observed that elasticity of substrate was increased

from 0.13MPa to 1MPa when the diameters and heights of nano-pillar were increased. On the

other hand, elasticity of 3T3 fibroblast varies with diameter and height of the Silicon substrate of

which the lowest diameter and height of pillars have highest cell stiffness. It was also found that

pillars with same diameter but with different heights have similar elasticity for cells and

substrates which suggests that different pillar heights do not have significant effect on elasticity.

Increasing the cell thickness the Young’s modulus decreased, indicates that leading edge have

higher stiffness than other part of the cell. Cells were well spread and firmly attached on the

Silicon nanostructure and movement of cell was taken place on the top of the pillars.

Chapter 1: Introduction Page 12 of 63

Chapter 1. INTRODUCTION

This chapter discusses the methods for fabrication of nanostructure substrates and the

relationship between the nanotechnology and biotechnology. In addition, the importance of

studying cell mechanics are explained.

MATERIALS SCIENCE AND ENGINEERING

Materials Science refers to the investigation of the relationship between the structure and

property of materials whereas materials engineering focused on the design and synthesis of the

new products or materials. Structure and property are the most important in materials science

and engineering, including processing and performance that influences the structure and

property of materials [1]. Nanomaterials exhibits some special chemical, mechanical, optical,

magnetic and electronic properties on surface due to the size of the material on micro to nano

scale. The optical property of nanomaterials varies a lot from the bulk material due to the

differences in the refractive index which measures the electromagnetic radiation. Similarly

mechanical properties such as strength and elastic modulus of bulk materials will be different for

nanomaterials [2].

1.1.1 Nanomaterials

Nanomaterial is regarded as the materials with a size range of 1-100 nm. Nanotechnology refers

to the process and synthesis techniques and characterisation of nanomaterials (crystalline and

amorphous) in detail. The synthesis or fabrication of nanomaterials is important due to its small

size and the mass with high surface area [3].

There are two conventional methods for fabrication, bottom-up and top-down. In top-down

approach, the bulk material are chopped, layer by layer, to a small material [4]. Consequently,

the waste is high but the method is fast and has very good control on particles shape and

spacing. Photo-lithography, Reactive Ion Etching (RIE), ball milling etc. are considered as the top-

down process [5]. On the other hand, the bottom-up process is similar to making a building by

placing bricks one by one. Nanostructure manufacturing or synthesizing is carried out by

assembling atoms or molecules. This process can be carried out from homogeneous nucleation

from liquid and vapour or heterogeneous nucleation on substrate, which is able to give properly

ordered nanostructure by means of building block. Wet chemistry routes e.g. precipitation,

reduction, sol-gel process, chemically and topologically pattern surface, organic block copolymer

are the most common types of bottom-up process to fabricate nanostructure [3].


1.1.2 Biomaterials

Biomaterial science mostly focuses on the interaction of materials in the biological environment

although the study is surrounded by physics, chemistry, biology, engineering and medicine.

‘Biomaterials’ as defined by DF Williams in 1987, ‘is a nonviable material used in a medical

device, intended to interact with biological system ’[6] . The material that should be used as

biomaterial should have biocompatibility and can perform specific tasks. The materials should

not be toxic, should be mechanically stable, should not make corrosion or degradation in vivo

and should be non-carcinogenic [6]. Nanomaterials can be used as biomaterials which can help

to develop new device such as diagnostic sensor or drug delivery system with precise dosage [7].

Fig 1-1 shows the relationship between the nanotechnology and biology and application of

them. The dashed line indicates that there is a possibility to make bionanodevice and

bionanosystem.

Figure 1-1: Correlation of nano and biotechnology [8]

Presently several materials including polymers have been used as biomaterials, for example,

polyurethane used in heart valve, teflon in vascular graft, hydrogel as contact lenses,

hydroxyapatite in healing bones, titanium alloy and some ceramics i.e. alumina used in dentistry,

polyethylene in hip prosthesis etc. [3].

Silicon (Si) as a material has wide range of applications from the kitchen to computer chips to

the human body. The use of Si as a biomaterial has been going on for decades due to

biocompatibility and biodurability and other chemical properties such as low surface tension,

ans hydrophobicity. Si provides one of the best biodurability, however, Si elastomers have lower

tensile strength or tear resistance than other elastomers. Si elastomers have been used in blood

coagulation prevention since 1940 because of its hydrophobicity. Some other important


applications of Si are in orthopaedics (hand and foot implants) kidney dialysis, blood oxygenator,

aesthetic implants such as breast implant etc. [6] [9].

IMPORTANCE OF STUDYING CELL MECHANICS IN BIOMATERIALS

In the biomaterials industry it is crucial to study the cell-materials mechanics due to interaction

of cells with different materials, such as in implanted devices where cells interact with materials

at nanoscale. The topography of material has significant influence on cell response [10] for

example, when the substrate has nano island on the surface, the cell morphology and focal

adhesion will be significantly different from the flat surface [11]. Furthermore, depending on the

materials that cells are interacting, cell response will vary. Cells will interact with metals

differently than polymer due to the high stiffness and stability and ordered atomic structure of

metals [12]. Cell mechanics has a great effect on cell proliferation, migration and differentiation.

It is also necessary to investigate the mechanical response of the cells to an external force such

as any chemical and physical signals. Heart is beating by expansion and contraction so blood is

pumped out to the body consequently, that creates a mechanical stress to the cells [13].

Additionally, analysing nanomechanics is getting important in cancer cell research. The variation

in elasticity in normal cell and metastatic cell can be measured in nanomechanics by using

Atomic Force Microscopy (AFM) [14]. There are other ways for quantitative analysis of

mechanics of cells such as Micropipette aspiration (MA), Magnetic Twisting Cytometry (MTC),

Optical and Magnetic tweezers. However, there are limitations using these techniques, for

example, in AFM the results rely on the spring constant of the tip of the cantilever and the

interaction between the tip and the cell surface so, there is a chance that the mechanical

property might be misinterpreted [15] [16].

In this study, fabrication of nano-pillars of Silicon was done using bottom up and top down

methods to investigate the mechanical properties of cells more specifically; elasticity of 3T3

fibroblast using AFM.

Chapter 2: Background Information Page 15 of 63

Chapter 2. BACKGROUND INFORMATION

This chapter includes detailed information of cell structure, functions and mechanics of the cells-

surface interaction. For the statistics the Hertz model has been used therefore, the theory of

elasticity and Hertz Model are also explained.

AN INTRODUCTION TO CELL

Cells are living organisms and the basic unit of life, and therefore perform and control many

body functions. There are two main types of cells: Eukaryotic and Prokaryotic. Animal cells

belong to Eukaryotic type. The component of a Eukaryotic cell are Nucleus, Cytoplasm,

Cytoskeleton, Golgi apparatus, Endoplasmic reticulum etc. [17]. Figure 2-1 shows the

components of a Eukaryotic cell.

Figure 2-1: Schematic diagram of a Eukaryotic cell [18]

The cell is surrounded by an Extracellular matrix (ECM), which is a non -cellular component with

strong biochemical and biomechanical behaviour and are responsible for tissue morphogenesis.

Two main classes of ECM are proteoglycans (PGs) and fibrous protein which are collagen,

fibronectin, elastin and laminins. These have different shapes and sizes with structural and

adhesive functions [19] [20] . The components of ECM are responsible for organizing the

orientation of the matrix such as cytoskeleton, which is situated inside of the cell and it’s

orientation can be controlled by the matrix situated outside the cell [20].


Fibroblast are situated in the loose connective tissues in ECM (Figure 2-2) and plays an important

role by proliferating, migrating and producing the collagen matrix whenever a tissue is injured

which helps damaged tissue to be repaired. However, the skin fibroblast cells are different from

others and they show different plasticity in the same cell culture [21].

Figure 2-2: Schematic diagram of Fibroblast in ECM [22]

Every eukaryotic cell possess an internal skeleton called the cytoskeleton. There are three main

components of cytoskeleton: actin filaments, microtubules and intermediate filaments. These

components are associated with proteins [23] and give the cell mechanical stability, shape and

capability to move from one place to another [17]. They form a network that inhibits any

deformation but when any force is applied form outside they can reorganize and maintain the

intracellular arrangement. The order of stiffness between three components are, microtubule>

actin filament> intermediate filament. Actin filaments are highly organised by proteins, and

possess isotropic, bundled and branched networks which are responsible for cell to cell

communication. Actin filaments and microtubules are associated with polarized subunit of

polymer, but intermediate filaments are not polarized and not able to assist with movement of

the cell [24]. Figure 2-3 shows the structure of the three components where, microtubules is the

biggest in diameter compare to others.


Figure 2-3: Schematic diagram of components of cytoskeleton, a) Microtubules b) Intermediate filament c) Actin filament [25]

The link between ECM and cells is maintained by Integrin, a cell adhesion molecule, and a

receptor protein. Integrin usually has two subunit of transmembrane glycoprotein called α and

β, which are non-covalently associated [26].

CELL –SURFACE INTERACTION

Fibroblast moves over the surface by crawling smoothly in cell culture and when it moves, it

makes elongated triangular formation. One of the triangle sides forms lamellipodia, but the

other two sides try to move backward or even remain motionless [27]. Movement involves three

steps, the leading edge extend and attach to the substratum then backside of the cell is pulled

forward, figure 2-4 [28].

Figure 2-4: Movement of cell by crawling over the surface [28]


Living cells feel different types of forces such as shear stress, compression and stretching and

hence have contractility which has an impact on cell functions. Wound healing, migration and

cytokinesis are regulated by the cell’s contractility in which the interaction mechanism between

the proteins of actin filament are responsible [29]. In figure 2-5 shows how the living cell binds

to the surface through integrin and can feel some forces.

Figure 2-5: Schematic representation of a cell possessing different types of forces [30]

When living cells are subjected to a substrate, they interact through transmembrane receptors

like integrin, a component of ECM on the substrate. Integrin forms a complex in the intracellular

side through focal adhesion and as a result ECM connects to the actin cytoskeleton or stress

fibres. Cells are also able to sense the substrate rigidity and therefore, adopt their structure and

can respond to forces as little as 5 pN [30].

THEORY OF ELASTICITY (HERTZ CONTACT MODEL)

Elasticity is defined as the regaining of the original formation when an applied forces is removed,

as the applied forces causes deformation of the structure. All materials possess elastic property

to a certain point [31]. Elasticity is described by Hooke’s law in terms of stress (𝛔) and strain (ε)

where, stress is an externally applied force per unit area and strain is the amount of deformation

caused due to force [1] [32].

According to the Hook’s law stress is proportional to strain in the form:

𝛔 = Eε (2.3.1)

E is a constant and it is called elastic modulus and dependent on the materials [32]. However,

elasticity of soft materials such as biological materials is determined by Hertz Contact Model

which depicts the elastic deformation of two homogeneous bodies contacting each other. This

model is widely used to measure the elastic property in a time scale by collecting force-distance

curve [33]. This model considers some assumptions such as:


The tip and cell material properties are homogeneous and isotropic;

The geometry of the communication between the tip and cell is axisymmetric, steady and

smooth;

The contact between two bodies is adhesionless and frictionless [34].

In this work spherical tip is used and the force (F) acting on the cell is described by

𝐹 =4

3𝐸∗𝑅

1

2 𝛿3

2 (2.3.2)

Here, E* is the effective Young’s modulus and R is the combined curvature of the tip (R1) and cell

(R2) and δ is the indentation depth. It also presumed that the cell is flat so its radius goes to

infinity. Since the tip is harder than the cell, the Young’s modulus of the tip is also considered to

be infinity [33]. For the force-distance curve and for fitting the curve the equation (2.3.1) will be

applied.

Figure 2-6: Schematic diagram of Hertz contact model with a spherical tip (radius R2), loading force F, Cell radius R1, a is contact radius and δ is indentation depth [34]

The effective Young’s modulus is express as [33]

𝐸∗ =𝐸

(1−𝜐2) (2.3.3)

Where, 𝞾 is the poisson’s ratio and for Hertz model the assumed value for biological sample is

0.5 [35].

Combining two equation (2.3.1 and 2.3.2), the expression of the Young’s modulus is [33]

𝐸 =3

4𝐹(1 − υ2)𝑅−1/2𝛿−3/2 (2.3.4)

It should mention that for the spherical indenter, the Hertz model will match satisfactorily if the

indentation depth is smaller than the curvature of the tip [36]. Although, the Hertz model is

lacking in giving absolute measurements it is still valuable to differentiate mechanical properties

between living cells and substrates.

Chapter 3: Experimental Page 20 of 63

Chapter 3. EXPERIMENTAL

This chapter discusses the materials and methods used in this study. This includes spin-Coating,

Dip-Coating, H2 Plasma, Photochemical growth and Reactive Ion Etching (RIE) for fabrication of

the nanostructure surface. Along with, the principle of the High Resolution Scanning Electron

Microscope (HRSEM) and Atomic Force Microscopy (AFM) explained here. An introduction to

Cleanroom technique also given. Cell culture preparation and calibration of AFM cantilever have

been explained.

CLEANROOM TECHNIQUE

Cleanroom is an environment clean from pollutants such as dust and chemical vapour.

Cleanroom technique controls temperature, humidity, pressure and electrostatic charge which

prevents contamination and maintain particulates i.e. oil, grease, hair etc. Cleanroom is required

to manufacture the healthcare products, pharmaceutical products as well as in industry research

on small particles [37].

The classification of cleanroom express as a number such as ‘class 100’, ‘class 1000’ and so on.

Classification depends upon the cleanliness of air in terms of particle diameter and density. For

instance, ‘Class 100’ means in each cubic foot there are less than 100 particles which are larger

than 0.5 microns and it is equal to ISO 5 cleanroom [38]. In this study fabrication was carried out

in cleanroom class 100.

Figure 3-1: Classification of cleanroom [39]


SPIN COATING: DEPOSITION BY SPINNING

Spin coating is a widely used technique to prepare a thin film on substrate in nanotechnology

because it is easy and fast and can be uniformly deposed on the substrate [40]. The precondition

for spin coating technique is the layer-building material should be viscous to float on the surface

and soluble in an evaporating solvent with an acceptable evaporating rate. it is also necessary

that the solvent has higher evaporating rate to get a homogeneous and failure free layer [5].

Coating the substrate should be on a turntable and the liquid should be placed on the middle of

the substrate. The rotation will start with a lower angular velocity which will be then rapidly

increased. The liquid solution then spread towards the substrate edge and consequently, a

uniform thin film will be formed [41] [42] [43]. Figure 3-2 shows different steps of Spin-Coating.

Figure 3-2: schematic diagram of different steps of spin coating: a) dispensation of photoresist, b) acceleration, c) spreading of the liquid, d) evaporation [40]

The viscosity and density of the liquid photoresist and the thickness and the rotation time are

main factors in spin coating. Higher spin speed and longer spin time form thinner film. The

interaction between solution and air is not as strong compared to the interaction between the

substrate and the solution. The thickness of the layer depends on the viscosity and

concentration of the fluid. When the solvent evaporates, the concentration of the fluid increase

hence the viscosity is increased [44] [42].

In this study, Si (100) n-type (phosphorus) (CrysTec, Germany) wafer was used as substrate with

a diameter of 3 inch and thickness of 380 µm. A thick positive photoresist, PMMA (poly methyl

methacrylate) which is a commercial e-beam/deep UV photoresist supplied by ‘Allresist’ has

been used on Si wafer to coat the whole surface by using ‘Convac 1001S’. This coating helped to

adhere broken particles which was generated during the cutting of the wafer by diamond tip.


At first, PPMA was spread on the wafer at low speed (rpm 500) for 10 seconds. Later the speed

was increased to 2000 rpm and ran for 40 seconds in order to dry the solution. The samples

were cut into the size of 10x5 mm2 to be analysed in High Resonance Scanning Electron

Microscope (HRSEM). After cutting, the substrates were cleaned in ultrasonic bath with acetone

and isopropanol for 12 minutes to remove the photoresist, and immediately dried in nitrogen

flow to avoid the film coating of acetone and isopropanol.

DIP COATING: PREPARATION OF NANOPARTICLES

Dip coating method is an easy and faster method for the preparation of the thin film from a

chemical solution. There are several ways to dip-coat such as drain Coating, angle-dependent dip

coating and classical dip coat from different types of solution. Solution could be inorganic

precursor or metallo-organic precursor and can be prepared by hydrolysis or condensation or

self-organization. Self-assembly principle, a spontaneous organization due to non-polar

interactions i.e. H-bond, Van Der Waals force, London force, is one of the best methods for

nano-structuring and thin film formation. The chemical solution contains amphiphilic molecules

that are composed of a hydrophilic and a hydrophobic part [45]. This process is usually based on

three separate steps as shown in figure 3-3.

Figure 3-3: Schematic diagram of stages of dip-coating process [46]

i) Immersion and dwell time: The samples are dipped into the precursor with a

constant velocity and defined time. Time is needed for the interaction between the substrate

and the coating solution to complete the wetting.

ii) Deposition and drainage: The substrate is then pulled out with a constant velocity.

When the substrate is pulling out and upwards there is a flux and excessive solution is drained


out. This specific velocity is important for determining the distance between the particles on the

substrate surface.

iii) Evaporation: The evaporation from the fluid is occurred and hence the thin film

deposition is made [45].

In this work, PS-b-P2VP (Polystyrene-b- poly-2- vinylpyridine) was used which is a diblock

copolymer and supplied by ‘Polymer Source, Inc’. This is a better approach for making

homogenous nanostructure array after which the copolymer can be removed completely. The

polymer is used with an appropriate non-polar solvent, toluene, to get the deposition of the

nanoparticles nearly ordered array on flat surface [47]. Poly(2-vinylpyridine)(P2VP) is hydrophilic

and forms the core while Polystyrene (PS) is hydrophobic and forms the outer shell. PS-b-P2V

polymer is dissolved in toluene, therefore spherical reverse micelles formed in the solution. With

the addition of salt of gold (HAuCl4) in the solution and stirring, gold migrates into the core of

the micelles, figure 3-4. Micelles are loaded with the same amount of metal salt at the

equilibrium [47] [48] [49].

Figure 3-4: Preparation of Au micellar solution [50]

PS(1800)-b-P2VP(770) was mixed with toluene and kept for one week under magnetic stirring.

Gold (lll) Chloride hydrate salt from ‘Sigma-Aldrich’ was then added and kept for another week

under magnetic stirring to help the polymer to be self-assembled. The structure of the PS-b-

P2VP and the preparation of micellar solution are given below.

Figure 3-5: Structure of PS-b-P2VP blocked copolymer, modified from [51]

Table 3-1 Preparation of micellar solution

Item Identity Quantity

Polymer(PS-b-P2VP) 1800-770 100 mg

Solvent Toluene 20 mL

Salt HAuCl4.H2O 50 mg


Before using the solution, filtration was done to avoid any potential contamination or pollution.

For filtration, three different types of filter media was used with different pore size, Table 3-2.

Table 3-2 Description of different types of filter

Filter Type (can be used in) Pore size

CHROMAFIL Xtra PTFE Non-polar media 0.45 µm

CHROMAFIL GF Highly contaminated media 1.0 µm

Millex FG Hydrophobic Flouropore 0.2 µm

After every dip coating the substrate was checked in the High Resolution Scanning Electron

Microscope (HRSEM) (Hitachi S5200 at 30 kV) to measure the distance as it varies with the

solution. Depending on the distance between the particles the withdrawal velocity was different.

The different velocity were 2.8, 3.6, 4.2 and 4.8mm/min. The interparticle distance was

calculated using ImageJ software.

H2 PLASMA

Plasma is increasingly used in semiconductor technology for removing carbon contamination,

native oxide layer, and also used in etching process [52]. There are different types of plasma i.e.

Oxygen, Hydrogen and Argon plasma which reacts with the deposited molecule on the surface,

break them down and convert into volatile compound. H2 plasma is gaseous and electrically

neutral which contains electron, ions, neutral atoms and molecules. Hydrogen has very small

molecular weight and energy therefore, sputtering is not possible but removing organic polymer

from the surface is possible [53].

When hydrogen plasma generated, it creates chemically active species and ions i.e. H* and H+

with low kinetic energy. The mechanisms are following [52]:

H2 + e → H + H+ +e (3.4.1)

H + e → H* + e (3.4.2)

H+ + H → H* + H+ (3.4.3)

H*, H+, H and e are hydrogen radical, hydrogen ion, hydrogen atom and electron respectively.

These all have strong role to remove the chemicals and oxide layer from the surface [53]. In this

study, ‘TePla 100-E’ was used to remove the polymer completely from the surface of the

substrate. To use this device, first vacuum was created in the chamber with pressure less than

0.05 mbar for plasma ignition. Then H2 gas was allowed to the quartz chamber for 15 min at 0.25


mbar for pre-treatment. Finally, H2 plasma was carried out for 90 min to remove the polymer.

The power was 160 W, the frequency was 2.46 GHz and the pressure was 0.8 mbar.

PHOTOCHEMICAL DEPOSITION

After dip-coating and then H2 Plasma treatment, the size of the gold naoparticles on the

substrate are quasi-hexagonally ordered and the size is small (average size is 9nm).Therefore, to

make a bigger size of gold particles, photochemical deposition process was used. In this process,

a solution was prepared by mixing gold salt (HAuCl4.H2O) with Phtalatester and irradiated under

UV light. The advantage of using Phtalatester is that it does not evaporate in UV light and also

absorbs in low spectral range. Exposure time is important due to the size of the growth of the

particles [54]. Figure 3-6 shows the effects during exposure.

Figure 3-6 Photochemical growth of Au particle [54]

The principle of this process is that when UV is exposed to the gold complex, gold salt absorb the

UV light and Cl- ion from the gold salt solution is oxidised by absorbing the energy and the

reduction of gold particles take place. Here is the reaction,

2AuIIICl-4 ℎ𝜐→ 2 AuIICl-3 +Cl2 (3.5.1)

2AuIICl-3 → AuICl-2 + AuCl-4 (3.5.2)

2AuICl-2 → Au0 + AuCl-3 +Cl- (3.5.3)

From the reaction it can be seen that this process involves two steps – first, the Au is gradually

reduced to Au atom and followed by agglomerate with existing gold particles on the substrate to

make small metal cluster[55] [56].

After preparing the solution the substrate has been exposed to UV light by seeding machine

from ‘Karl SUSS MJB 3 Mask UV 400’, West Germany. Figure 3-7 shows the schematic diagram of

this seeding machine. This machine has an Hg (Mercury) short-arc lamp surrounded by the

ellipsoidal mirror and maximum power (350 W) of the light can be used at two wavelengths:


365nm and 390nm. When the radiation was discharged by turning on the light, the radiation is

collected by the ellipsoidal mirror and then focused to the cold light mirror. Only short

wavelength light is reflected to Fly’s eye lens and the condenser lens adjust the light intensity.

There is a filter to block undesirable wavelength, but in this work no filter was used. By using the

lens plates it is possible to remove diffraction effect during the experiment. With the help of

surface mirror and front lens, the beam exposes to the substrate vertically [57].

Figure 3-7: Schematic diagram of optical system of mask aligner Karl SUSS MJB 3 Mask UV 400 [58]

After putting the substrate on the chuck, 20µl of the solution has been poured on each substrate

using a pipette. Exposure time was 1.5 minutes for 17nm diameter and 3.5 minutes for 30nm

diameter of the particles. Then the substrates were cleaned with acetone and isopropanol for

12min and dried in nitrogen flow. But there is a chance that the organic molecule could be

present on the substrate so H2 plasma was carried out again for 15min. The preparation of gold

salt solution is given in Table 3-3.

Table 3-3 Preparation of the gold solution for photochemical growth

Phtalatester Gold Salt (HAuCl4.H2O)

Density 1059 g/l Molar mass 339.79 g/mol

Weight 2056.99 mg Weight 3.3 mg

Volume 1.95 ml Concentration 0.005 molarity

Plasma Etching: Reactive Ion Etching (RIE)

Etching is the process of removal of materials from a substrate. There are two main division of

etching: wet chemical and physical dry etching. In wet chemical, materials are removed by using

liquid chemicals or etchants. Specific patterns are protected by masks, otherwise the whole

surface will be etched away by liquid chemicals. In wet chemical etching there are three basic

steps are followed [59]:


The liquid etchant diffused to the structure;

A redox reaction occurs between the liquid etchant and the materials to be removed;

By-products diffused from the surface

On the other hand, the physical dry etching process is carried out in gaseous phase by high

kinetic energy such as particle bombardment, chemical reaction or combination of both

followed by evaporation. In general, when the high kinetic energy ion, electron or proton in

touch the substrate knocks out the atoms from the surface. There are three different types of

dry etching: Chemical Plasma Etching (PE), Reactive Ion Etching (RIE) and Ion Beam Etching (IBE).

RIE gives high etch rate and high selectivity due to combination of physical sputtering and

chemical activity [59][60].

RIE is plasma assisted etching which involves the generation of glow discharge of a feed gas, i.e.

CF4 for Si etching, by which high kinetic energy particles, neutral atoms, electrons, radicals and

positive/negatives ions are produced. Since the substrate is placed on the coupled electrodes

and obtains a negative charge so the positive ions are attracted to the substrate and diffused to

the surface to start etching [61]. Figure 3-8 shows the schematic diagram of this process. This

process is associated with several steps, such as formation of ions, radicals, diffusion,

adsorption, chemical reaction desorption and pumping out the reacted product.

Figure 3-8: Schematic representation of the process of etching adopted from [61]

Bombardment makes free radicals as active sites which then adsorb and react with the

substrate. For example, Fluorine bombardment in Si wafer etching, reaction between F atom

and Si produces SiF4, which is volatile. To desorb the SiF4, a high vapour pressure is needed at

substrate temperature. Reacted products mostly go back to the plasma region therefore, it is


necessary to pump out afterwards otherwise, there is a chance to dissociate with others and

resorption will take place [61].

In this work, ‘Oxford Plasmalab 80Plus ICP65’ has been used for the RIE and the etching process

occurs vertically therefore, anisotropic etching profile can be reached. The gases produced are

CF4, CHF3, O2, Ar, hence, the influence of these gases will be explained here.

Using CF4 gas gives the following mechanism [62]:

CF4 → F* + CF3 (3.6.1)

CF4 + e → CF3+ + F* (3.6.2)

Si + 4F* → SiF4 ↑ (3.6.3)

3.6.1 Effects of Oxygen addition

The mixture of different gases have different influence on etching rate. For instance, addition of

O2 (< 5%) to CF4 plasma increases the density of F atom and consequently rate of etching as seen

in following reaction. However, addition of O2 over 15% decrease the density of fluorine [63].

CF4 → F* + CF3 (3.6.4)

CF4 + e → CF3+ + F* +2e (3.6.5)

O2 → O* + O (3.6.6)

O2 + CF*x → CO2 + COF2 (3.6.7)

CF4 + O → COF2 (3.6.8)

3.6.2 Effects of Hydrogen addition

The addition of H2 on CF4 plasma reduces the density of F atom thus etching rate due to the

formation of HF. H2 also reacts with CF3 radical and produce polymeric precursor which forms a

layer of CxFy on the surface. At high concentration of H2 (> 30%) etching will stop due to the

polymerization on the surface [60].

3.6.3 Effects of CHF3 and Noble gas addition

Addition of CHF3 in CF4 plasma does not make any significant changes because CF4/CHF3 is very

similar to CF4/H2 system, however, CHF3/HF has lower internal energy than CF4/H2 system.

CHF3/HF can be produced by mixing CF4 with H2 [60].

Noble gases, mostly Argon (Ar) and Helium (He) are added to stabilize the plasma. Addition of Ar

can make ion bombardment on the surface consequently, it increases the anisotropic etching.

Helium is used in order to cool the substrate from the front or back side [60].


Finally, the Au NPs were removed by Loguls solution and then rinsed with Millipore water.

Loguls solution was prepared by adding 4gram of Potassium Iodide from ‘Prolabo’, 1gram of

Iodine from ‘Merck’ in 150ml deionised water.

ELECTRON MICROSCOPE

To study the structure of the feature in nanometer range, Scanning Electron Microscope (SEM)

and Atomic Force Microscope (AFM) have been used. Furthermore, using electron microscope, it

is possible to see the single atom and soft materials like biological cells in a very low voltage.

3.7.1 Scanning Electron Microscopy (SEM)

SEM gives the information about the topography, morphology, composition and crystallographic

structure of the feature. A voltage is applied to the electron gun to heat up the filament

(cathode) which then emits thermo-electrons after it reaches a defined temperature [64]. The

filament is usually made of tungsten which is about 0.1mm and is heated up to about 2800K. The

thermo-electrons known as electron beam, are then forced to go to anode by applying positive

voltage (1-30kV) to the anode [65], figure 3-9.

Figure 3-9: Schematic diagram of Scanning Electron Microscope (SEM) [66]

When the high energy electrons enter to the specimen they scatter and lose their energy. Some

of them are absorbed in the specimen, and some of them are emitted from the specimen as

secondary electrons, back scattered electrons, Auger electrons, figure 3-10 [65]. Secondary

electrons are produced from inelastic collision with the atom of the specimen, and with an

energy less than 50eV provides information about the topography of the specimen. The


backscattered electrons produced from elastic collision are higher energy greater than 50eV

[67]. These electrons come from a deeper region of the specimen and are sensitive to

composition, therefore provides information about the atomic number [65]. In this work, Hitachi

S5200 High Resolution Scanning Electron Microscopy (HRSEM) was used with a cold emission

gun and tungsten cathode.

Figure 3-10: Emission of various electrons and electromagnetic waves from the specimen [65]

3.7.2 Atomic Force Microscopy (AFM)

AFM is one of the best modern techniques in biomaterials and nanomaterials field and has a

great contribution in cell study. The working principle is simple comparing to electron

microscopes; AFM detects the forces acting between the AFM tip (attached to a very flexible

cantilever) and the surface of the substrates [68]. Figure 3-11 shows schematic diagram of an

AFM setup where a laser light is focused on the back side of the tip, which is reflected and then

detected by the photodiode. To form an image, cantilever comes close to the surface of the

sample and scans line-by-line. By doing so it feels deflection due to the tip-sample interaction.

This deflection is detected by the photodiode detector [69].


Figure 3-11: Schematic diagram of AFM [69]

AFM possesses a piezoelectric scanner that moves over the surface of the sample. During image

acquisition, the scanner moves fast along the horizontal line and slow along the vertical line and

takes data points. The space between the data points is called ‘step size’ and for this experiment

64 data points were taken. Once it finished scanning across the horizontal line it comes back to

its perpendicular position and start scanning the second line and continues, figure 3-12 [70].

Figure 3-12: Schematic diagram of scanning system in AFM, redrawn from [70]

In this work force spectroscopy mode was used in AFM which assists to measure force at a

specific point. Here, the tip and cantilever move up and down to the surface. When the distance

between the tip and surface of the sample is big, no deflection is recorded (Force=0) [68]. When

tip-sample distance decrease (approaching red colour in Figure 3-13), at some point tip jumps

into contact to the surface (attractive force) and this effect called ‘snap-in’. When the cantilever

contacts with surface it applies some forces, from here it is possible to investigate Young’s


modulus or stiffness of the surface. After that at some point cantilever feels repulsive force and

retracts (blue colour in figure 3-13) from the surface but for a while tip tries to keep in contact

because of adhesion [71].

Figure 3-13: Force spectroscopy mode in AFM [71]

For this experiment, AFM NanoWizard 3 (JPK instrument, Berlin, Germany) was used and there is

an optical microscope (Zeiss Axiovert 200) fitted to see the cells. The cantilever is made of

Silicone (B500-CONTAuD-5), coated with gold. The tip is spherical and high density diamond-like

carbon with 500 nm ±10% in diameter. The nominal spring constant is 0.2 N/m, but actual spring

constant derived from the calibration data.

Before running the experiment calibration of the cantilever was carried out at its resonance

frequency by thermal noise method which is commonly used and highly automated [72]. The

spring constant was determined in two steps. First, from the slope of the force curve which

shows the sensitivity of the cantilever and second, resonance frequency from the spectrum [34].

JPK software was used in contact mode on a hard surface i.e. glass in liquid, in this case DMEM

medium so, that there will be no indentation of the surface for the calibration. Figure 3-14

shows the spectrum of the fluctuations of the cantilever as a function of frequency, and from

thermal noise data the value of the spring constant was calculated to be 0.147N/m [71].


Figure 3-14: Calibration of cantilever spring constant, blue line is for cantilever resonance peak

(measured) and red is the Lorentz fit [71]

For imaging, quantitative imaging (QI) mode was used, which is developed for AFM by JPK

instrument, which works while not applying lateral force therefore, it helps to control the

vertical force at every pixel. This mode makes AFM imaging easy and faster by controlling tip-

sample force at each point of the image. There are other advantages of using this mode, such as,

no need to adjust the set point or gain during scanning and it also gives the information about

elasticity, adhesion and dissipation [73].

The settings of the AFM are given below:

Table 3-4: Settings of AFM

Setpoint 4nN

Z length 3500nm

Extend time 100ms

Extend speed 35µm/s

Retract time 35ms

Retract speed 100µm/s

Fast 30µm

Slow 30µm

X-Offset 0µm

Y-Offset 0µm

Grid angle 0 degree

Pixels 64*64

Pixel ratio 1:1


Extend sample rate 100kHz

Retract sample rate 100kHz

Add. retract 50nm

Motion time 5ms

Acceleration 1.5ms

Time for image 9.73min

CELL CULTURE PREPARATION

In this work 3T3 fibroblast was used which is an embryonic mice skin cell supplied by Medicine

Department in University Ulm. First, the substrate was sterilised with ethanol and dried at room

temperature. Second, fibronectin coating was made on the Si substrate to adhere fibroblast on

the surface. For fibronectin coating, 5% solution was poured on the substrate for 2 hours. After

preparing the cells, they were placed into the incubator. The environment of the incubator is 37°

C, 98% humidity and 5% CO2 and kept for 24 hours. Cell culture was done by Ulla Nolte in

Experimental Physics department, University Of Ulm. The petri dishes uses for cell culture are

from ‘TPP’ made in Switzerland. These Si substrates can be reused for cell culturing once the

experiment is done.

Chapter 4: Results and Discussion Page 35 of 63

Chapter 4. RESULTS AND DISCUSSION

FABRICATION OF NANO-PILLARS

One of the part of this thesis work is to fabricate well-ordered nano-pillars, and this involved

several steps. The results of these steps are given here.

4.1.1 Micellar technique by using block copolymer

Micellar technique is comparatively easier way to make thin film on a substrate. The distance

between particles can be determined by pulling out the substrate with a constant velocity, figure

4-1 or changing the concentration of the solution, figure 4-2. In this work, the substrate was

pulled out with a constant velocity to determine the distance of particles.

During dip-coating, the substrate was withdrawn vertically with a constant velocity U. The model

for dip-coating given by Landau and Levich is:

h = 0.946 * √𝜎

𝜌𝑔 Ca2/3 (4.1)

Here, h is the thickness of the wetting film on the substrate after removal from the micellar

solution, Ca is the capillary number, is surface tension and 𝜌 density of the solvent.

Now, equation (4.1) will be valid when the capillary number, 𝐶𝑎 = µ𝑈

σ, where µ is the dynamic

viscosity [74]. The maximum thickness will be proportional to U2/3. After evaporation of the

solvent, the thickness is proportional to the deposited micelles and the areal density of the

deposited micelle, monolayer film is also proportional to U2/3 [75]. Therefore, the interparticle

distance is proportional to U-1/3 [58]. From Figure 4-1 it is observed that when the velocity

increases the distance between the particles decreases and vice versa. In addition, with constant

interparticle distance the pulling out velocity varies depending on the types of substrate. For

example, if the distance was kept at 100nm, the velocity for Si was 4.8mm/min while for SiO2 the

velocity was 10mm/min.


Figure 4-1: Plot of average interparticle distance over withdrawal velocity from experimental data, from figure it is seen that for 130nm interparticle distance the withdrawal speed is 2.8mm/min

Figure 4-2: Interparticle distance changes depending on the concentration of the solution [58]

After dip-coating, polymer was removed from the surface through H2 plasma so that the

distance between the Au particles can be measured. The Au NPs are arranged hexagonally

which also indicated the particles were self-assembled on the substrate. Figure 4-3A shows the

hexagonal distribution of particles taken by HRSEM at 30kV while figure 4-3B shows the

threshold image using ImageJ software 1.46r version by applying bandpass filter. The inset of

figure 4-3B shows the hexagonal arrangement of Au particles. The average distance between the

particles was measured at 130nm.

Figure 4-3: HRSEM image of Au nanoparticle after H2 Plasma. A) Grey scale picture and B) after adjusting threshold, inset hexagonal arrangement (bandpass filter). Scale is 1µm, 30kV.


4.1.2 Photochemical growth of Gold (Au) particle

Nanoparticles have size dependent properties, therefore, the precise fabrication is important.

Photochemical growth is one of the easiest and fastest method to seed particle [76]. It is

important to mention that depending on the number of monomer of the block copolymer, the

size of the gold particles may vary. It has been shown that by using PS(325)-b-P2VP(75), the size

of the Au NPs was 2.9±0.4nm while using PS(1350)-b-P2VP(400), the size of Au NPs was

7.9±1.2nm [47]. In this work PS(1800)-b-P2VP(770) was used as block copolymer and the

average diameter of the gold particle was found to be 9nm. The photochemical growth process

was conducted with exposure time 1.5 minutes and 3.5 minutes and found that the average

diameter of the Au NPs are 17nm and 30nm respectively. Figure 4-4 shows the relation between

diameter and exposure time.

Figure 4-4: Plot of Au NPs diameter as a function of exposure time from experimental data.

Figure 4-5 shows that particles were getting bigger after seeding. In general, particles up to

30nm are monodisperse and longer reaction time will cause the dislocation or disorder of the

particles [76]. After dip-coating the particles were found to be spherical but when the size was

increased the particles were getting less and less spherical. The reason could be that particles

were contaminated with air while transporting the substrate to the cleaning box or the humidity

was not ideal.

Figure 4-5: HRSEM images (at 30kV) of different diameter(average) of Au NPs, a) 9nm, b) 17nm, c) 30nm, scale is 200nm.

a)

b)

a)

c)

a)


Less spherical particles can be made spherical by annealing at 720°C for 1 hour afterwards [77].

The diameter of Au NPs were determined by converting the picture to its threshold then

analysed the particle area of each particle in the software as seen in figure 4-6.

Figure 4-6: a) grey scale image using HRSEM scale 200nm at 30kV; b) after adjusting threshold; and c) marking for area measurement through imageJ software

4.1.3 Reactive Ion Etching (RIE)

CF4 gas is used to etch to fabricate well-ordered nano-pillar array using Au NPs as mask in

Reactive Ion Etching (RIE). Although CF4 gas yields higher etching rate but it also removes Au

nanoparticles. Therefore, a fluorocarbon layer close to the Au NPs is necessary to reduce under

etching [78]. During the experiment, the mixture of CF4-CHF3 gases with flow rate 2sccm:20sccm

(Standard Cubic centimetres) and low pressure 1mTorr was used. The temperature was

maintained at 25°C by liquid nitrogen. The DC bias was maintained at 96V by changing plasma

power between 56 and 63W, and the system was operated by PC 2000 software.

In CF4 based Si etching, F radicals adsorb on the surface and react to produce SiFx layer on the

surface. Two F atoms form SiF2 on the upper level and are removed. But when more F atoms are

available, it forms SiF4 and desorbs [79]. Figure 4-7 shows the formation of SiF2 and SiF4.

Figure 4-7: Schematic diagram of a) formation of SiF2 and b) formation of SiF4 [79]

Depending on etching time the height of the pillars varies and therefore etching rate can be

measured by plotting height vs etching time. During etching time the rate and the shape of the

a)

b)

a)

c)

b)

a)

a) b)


pillar depends on DC bias. In the previous study Si etching rate was found to be 4nm/min with

30W power [78] while in this study the etching rate was 5.20nm/min with 56 to 63 W. Using high

power makes the applied voltage high and this makes ion bombardment faster hence, increases

etching rate. In addition, the aspect ratio (width over height) was experimentally found to be

0.52. A study showed that increasing power also causes decreasing selectivity [80]. Selectivity is

defined by the ratio of etching rate of two different materials. During RIE, erosion of gold atoms

occurs due to the sputtering. Therefore, the selectivity of these two materials was calculated to

be around 5. Another study [77] showed that depending on the size of the mask etching rate

varies, figure 4-8 but in this work it was not so obvious.

Figure 4-8: Etching rate changes depending on a) DC bias and b) size of the mask [77]

Figure 4-9: Au NPs in 200nm scale, tilted by 30 degree a) average height 75nm and diameter 28nm with Au particle diameter 17nm, b) height 75nm and diameter 49nm with Au particle diameter 30nm, and c) height 106nm and diameter 49nm with Au particle diameter 30nm

The diameter of the pillars was measured on top and on full width high maxima. For 17nm

diameter of Au NPs, the average diameter of the pillars on the top was 28nm and at FWHM

calculated to be 34nm. For 30nm Au NPs the diameter on the top was 49nm and at FWHM was

52nm. Since cells are interacting with pillars only on the top therefore, the top diameter was

counted for the measurement.

Table 4-1 shows the heights of the pillars with corresponding Au NPs diameter. It is observed

that for 17nm and 27nm diameter of Au NPs, the average pillar heights are a bit larger. It can be

explained that the DC bias was a bit higher (98 V) and therefore the rate was higher.

a)

))

b) c)


Table 4-1: Experimental data of particle diameter, etching time and average height

Average Diameter (nm) Etching time (min) Average height of nano-pillars (nm)

15 13 67

15 10 47

17 13 75

27 12 66

27 20 116

30 22 106

Figure 4-10 shows the AFM pictures using tapping mode. Some areas on the surface shows no

pillars due to scratches on the substrate surface during handling. The etching rate without any

mask was higher than with mask therefore, the pillars height adjacent to the scratch are higher

than the normal height of the pillars. Figure 4-10 (right) showed how the height was measured

by Gwydion software by making cross section over the scratch (yellow and red line). From Figure

4-11 it is seen that the height adjacent to the scratch was around 400-500nm and the scratch

area is rough.

Figure 4-10: AFM pictures (topography) of Samples 2 (130-28-75)


Figure 4-11: Height of pillars build by Gwydion, profile 1 corresponds to red line and profile 2 corresponds to yellow line from Figure 4-10 right part.

CELL MECHANICS ON NANOSTRUCTURE TOPOGRAPHY

This section discusses indentation depth, evaluation of the measurements, and comparison of

stiffness of cells and between the substrates. Cell-surface interaction are also discusses here.

4.2.1 Indentation depth

Hertz model is used for the measurements and there are two conditions for indenter. If these

two are met then Young’s modulus can be calculated from Hertz model. The conditions are that

the indenter will not deform and there will be no added collaboration between sample and

indenter. Figure 4-12 (top) shows the schematic diagram of indentation test, where the

cantilever is moved down to the sample by distance z, called ‘height measured’. But the

cantilever is bending in the opposite direction (x), indenting the sample by δ. The indentation

can be estimated by deducting the deflection of the cantilever from height measured. The

bottom figure is the deflection of the cantilever [14].

Figure 4-12: Schematic diagram of indentation test (top) and force-indentation curve (bottom) [14]


Finding the right indentation depth is important, otherwise measurement will vary. Although,

due to the properties of living cells such as elasticity, viscosity and adhesion, the elucidation of

the experimental data would be difficult. Figure 4-13 was drawn using Python and a program

named ‘hertzfit’ written by Christian Bühler [81]. Using this program it is possible to see

topography of one cell with Force-Distance curve. Figure 4-13C shows the contact point in the

force-distance curve (red point). If the hertzfit fits this point nicely then measurement of the

indentation depth can be done from figure 4-13B and 4-13D. Figure 4-13A shows the topography

of the cell, 4-13C is force-distance curve, 4-13B is ‘Young’s modulus error’ vs ‘indentation depth’

and 4-13D is ‘Young’s modulus ‘vs ‘Indentation depth’ curve. To get the acceptable indentation

depth, several force-distance curve with different fitting and different cells were investigated. In

figure 4-13 an indentation depth 100nm was chosen because at this point Young’s modulus start

to be steady. Once the indentation depth is chosen, the evaluation of all cells was carried out by

an extension of the program called ‘Hertzfolder’.

It is important to mention that if the indenter is on the nucleus the indentation depth will be

higher than if the indenter is far from the nucleus and importantly, small indentation depth (5-

10% height of the cell) is acceptable in Hertz model [14].

Figure 4-13: Sample 1: 130-49-106, Hertzfit indentation depth 100nm, figure A shows the topography of the cell, C is force-distance curve, B is ‘Young’s modulus error’ vs ‘indentation depth’ and D is ‘Young’s modulus ‘vs ‘Indentation depth’ curve

4.2.2 Measurements

Evaluation of data was done by MATLAB, scripts were written by Dr. Tobias Paust, Jonas Pfeil

and Fabian Port, University Ulm. For this work some changes have been made in the scripts. The

axis of the Young’s modulus was presented as the logarithm for a better view. The set point was

C D


4nN. For the experiment, it was tried to collect the data far from the nucleus which is only

cytoskeleton and substrate. Three different types of samples of Silicon has been used to

investigate the elasticity. Samples differ by their diameter and height. Samples are named by the

distance-diameter-height, all are in nm range. For example, Sample 1: 130-49-106, Sample 2:

130-28-75, Sample 3a: 130-49-75 and Sample 3b: 130-49-75.

4.2.2.1 Sample 1: 130-49-106

Figure 4-14 shows the histogram of the substrate with 3T3 fibroblast, which indicates that

elasticity distribution of the cell has a wide range of 102Pa to 104Pa. In contrast, for the substrate

the distribution is not wide. For this sample, the most frequent Young’s modulus for cells is, Ecell

~ 3.7kPa and for the substrate, ESubstrate ~ 1MPa. Different colours were produced by addition of

one cell with another with a total of 10 cells. The reason for wide range of elasticity value for

cells could be due to the different component of cytoskeleton. As stated in Section 2.1, the

stiffest components of the cytoskeleton is the microtubules so the value of Young’s modulus

more than 10kPa may represent the microtubules. Young’s modulus is between 1 and 10kPa

represent actin filaments. And the lowest value, less than 1kPa is for intermediate filaments. To

find out the contribution of these three components to the mechanics, immunofluorescence

imaging can be performed by using confocal laser scanning microscope.

Figure 4-14: Histogram of Young’s modulus by using Hertz model of 3T3 fibroblast of Sample 1 at indentation depth 100nm. X-axis is in logarithm, Y-axis is linear scale (calibrated spring constant 0.275 N/m), different colours produced by the addition of one cell with other and this results come from summation of 10 cells data.

A study [82] showed that 3T3 fibroblast has an elasticity range from 4-100kPa and authors

suggested that elasticity of cells comes from large contribution of actin filament rather than

other components. Other components such as intermediate filament also contribute but

microtubules do not have significant contribution to stiffness. Therefore, it can be said that in


these experiments most frequent elasticity belongs to actin filaments and the value more than

104Pa correspond to microtubules.

4.2.2.2 Sample 2: 130-28-75

Figure 4-15 shows the histogram of Sample 2, in this case the indentation depth was 100nm (30

cells measurements).

Figure 4-15: Histogram of Sample 2 at indentation depth 100nm 30 cells measurement, (calibrated spring constant-0.125N/m), different colours arise by adding one cell with other

In contrast with Sample 1 figure 4-14, the Young’s modulus of this substrate is less, which is

ESubstrate ~ 0.5MPa. From Figure 4-15, the distribution of the cell is wide between 102 and

5*104Pa. The most frequent elastic modulus of the cells measured in this case is Ecell~ 10kPa. It

was observed that decreasing the pillar diameter and height resulted in decreasing stiffness for

the substrate.

A study showed that the cell stiffness changes with the substrate stiffness and on a rigid surface

the cell spread well and the stiffness of the cell increased. Authors worked with human

mesenchymal stem cells (hMSCs) and found that with increasing substrate stiffness of 1-30kPa,

cell stiffness increased 1-7kPa [83]. In contrast figure 4-16 showed that elasticity of substrate

was decreased but the cell elasticity increased compared to the Sample 1. In Sample 2 Pillar

height and diameter are lower than Sample 1 and it also showed decreasing elasticity of the

substrate but increasing cell elasticity.

A study [84] mentioned that the Rotsch et al. (1999) investigated elastic modulus of 3T3

fibroblast and found that while the cortical stiffness for stable edge was 12kPa but for the

leading edge the stiffness was 3-4kPa. Comparing to the results in this study it can be reasonably

claimed that the stiffest part of the cell belongs to the stable edge of the cell due to the stress

fibres. Mahaffy et al. [84] investigated Young modulus by applying two different models, well-


adhered and non-adhered regions for 3T3 fibroblast, for the former one elasticity was

1.6±0.2kPa and for later, elasticity was 0.6±0.1kPa. Both are smaller than this work.

4.2.2.3 Sample 3a: 130-49-75

From figure 4-16 it’s observed that for Sample 3a, the most frequent elastic modulus for cell is

ECell~1kPa and for the substrate ESubstrate ~1.3*105Pa. The calibrated spring constant was

0.084N/m therefore, it can be said that with a soft cantilever the elasticity will be less.

Figure 4-16: Histogram of sample 3a at indentation depth 100nm, with spring constant 0.084 N/m, different colours arise from summation of all cells (6cells measurement).

4.2.2.4 Sample 3b: 130-49-75

In contrast with Sample 3a, Sample 3b (figure 4-17) has a stiffer spring constant (0.276N/m).

Therefore, the elasticity is higher than Sample 3a. Here, the most frequent elasticity calculated

for cell was ECell~4.3kPa and for the substrate was ESubstarte~1MPa.

Figure 4-17: Histogram of Sample 3b at indentation depth 100nm (5cells measurement) with calibrated spring constant 0.275 N/m, different colours arise from summation of all cells.

Since Sample 1 and Sample 3b have same diameter with different height of the pillars, the

results were compared to see the contribution of height on elasticity.


Figure 4-18: Histogram of Sample 1 (top, 10 cells) and Sample 3b (bottom, 5 cells), frequency scale is different due to the different number of cells measurements, calibrated spring constant 0.275 N/m. Different colours produced by the addition of one cell with other.

From figure 4-18 it can be observed that sample 1 and sample 3b (with same k and diameter of

the pillars) do not have significant difference in elasticity even though they have different height.

Therefore, it can be suggested that different height of the pillars do not have a significant effect

on the elasticity. The most frequent elasticity for the substrate were same but for cells, Young’s

modulus was Ecell~3.7kPa for Sample 1 and was Ecell~4.3kPa for Sample 3b.

To make comparison of Young’s modulus among the samples, boxplots were created, figure 4-19

showing the median at indentation depth 100nm. The corresponding median shows upper part

of the box. The highest median corresponds to Sample 3b, 6.02kPa and the lowest is 1.2kPa for

Sample 3a. The highest maximal belongs to Sample 2. Since the cell number are not same for all

samples eventually, the comparative results may not be fully representatives of the samples and

may need further study. Codan et al. [85], measured elasticity of living 3T3 fibroblast on glass

and found the Young’s modulus median is 5.2kPa. Codan et. al., used squared pyramidal tip

therefore the model is different from this study. Depending on the model, the elasticity values

will differ from each other.


Figure 4-19: Boxplot for comparing the median of different samples at indentation depth-100nm

Figure 4-20, Young’s modulus was plotted over height for the samples. It demontrates the

Young’s modulus of corresponding contact point through MATLAB Program. From here it is seen

that the Young’s modulus is decreasing with increasing height of the cell except Sample 3a,

which shows very different intervals compared to other samples. The highest Young’s modulus

was measured at near to zero micron. It should be noted that there could be 5-10% error in

measurement of height due to glueing the substrate onto the petri dishes. In the region of 2.5 to

3.5µm, the elasticity decreased for Sample 3a, but increased for other 3 samples and showed

very rough intervals. It can be explained that there are some area where cells overlaped on each

other and this height is coming from this overlaping. However, over 3.5µm height, the nucleus

could be the responsible for this height and shows very different elasticity under force. It was

mentioned before that the Hertz model is only valid at low indentation depth, but when the

indenter is on the nucleus area the indentation will be large. This may also explain this uneven

elasticity. The change of Young’s modulus with height also indicates that cells are

heterogeneous.

Median:

3843.8 Pa 4987.7 Pa 1193.3 Pa 6015.6 Pa


Figure 4-20: All samples: Sample 1: 130-49-106, Sample 2: 130-28-75, Sample 3a: 130-49-75 and Sample 3b:130-49-75

Solon et. al. [86] showed that the stiffness of 3T3 fibroblast differ depending on the distal

(17kPa) and proximal (5kPa) regions and higher parts of the cell are soft and homogeneous. In

similar to this study, authors found that the lower part of the cell is the stiffest and the

contribution of the substrate stiffness on cell stiffness. But authors found that over 700 nm of

height of the cell the Young’s modulus remain constant which is dissimilar with this thesis work.

It was suggested that when the cell thickness is more than 700 nm (same as this work), the

deformation of the cell distribute only into the cell body but were not transmitted to the

substrate.

Since Sample 3a shows rough intervals, to see details of Young’s modulus over height, boxplot of

selected heights such as 0.4-0.6 µm, 0.9-1.1 µm, 1.4-1.6 µm and 1.9-2.1 µm is presented in

figure 4-21. It is observed that the highest median correspond to Sample 3a in 0.4-0.6 µm but in

other parts it’s different. From here it is clear that something is wrong with Sample 3a. The

problem could be related to calibrated data.


Figure 4-21: Boxplot of all samples of Young’s modulus over selected height: 0.4-0.6, 0.9-1.1, 1.4-1.6 and 1.9-2.1 µm

From figure 4-21 it is seen that when the cell thickness is low the elasticity is high. Therefore, the

0.4-0.6 µm height the Young’s modulus is higher than others. But Park et. al. [87], showed

different result with 3T3 fibroblast, authors showed that elasticity increase with increasing cell

thickness (570nm-4700nm). The leading edge has lower elastic constant than the cell body

which is dissimilar to the current work.

Sample 3a was omitted from Young’s modulus over height plot and corresponding boxplot,

presented in Figure 4-22 and Figure 4-23 respectively.

Figure 4-22: Young’s modulus VS Height plot of 3 samples: Sample1: 130-49-106, Sample2: 130-28-75, and Sample3b: 130-49-75

Rotsch et. al., [88] worked with 3T3 fibroblast and showed that the dynamics is different for

active edge and stable edge. The leading edge height profile was rather flat between 0.4-0.6 µm


compared to the stable edge (2-3 µm) and the leading edge is softer than the stable edge. The

result contrasted with this study, increasing thickness of the cell, elasticity decreased and elastic

modulus was highest at the flat part, figure 4-22. Haga et. al., [82] discovered that nucleus has

10 times lower elasticity than its surrounding area. It is considered that over 3.5 µm height is

attributed the nucleus and it also shows lower elasticity than its surrounding area.

Figure 4-23: Boxplot of 3 samples of Young’s modulus over selected height, the selected heights are: 0.4-0.6, 0.9-1.1, 1.4-1.6 and 1.9-2.1 µm

From figure 4-23 it is seen that highest Young’s modulus corresponds to Sample 2 in all region. It

indicates that cells are stiffest on Sample 2 which has lowest diameter and height of the pillar.

On the other hand, Sample 1 and 3b have significant different elasticity with increasing cell

thickness. When thickness of the cell is low (0.4-0.6 and 0.9-1.1 µm), Sample 3b has higher

elasticity than Sample 1 but in thicker area (1.4-1.6 and 1.9-2.1 µm) Sample 1 has higher

elasticity than Sample 3b, indicates that cells stiffness is slightly dependent on pillars height with

increasing thickness of the cell.

It is calculated that in ~1µm2 area the average number of pillars in all samples remained same

but the average top area of the pillars that the cells were interacting with was different. In

Sample 2 it was 666nm2 but for Sample 1 and 3b the average area on the top of the pillars is

almost 3 times larger at 2078nm2. Therefore, Sample 1 and 3b should give the stability to the

cells better than Sample 2, but here it shows different. It should be noted that other samples

showed an indentation depth of 150nm as well but Sample 2 showed only 100nm indentation

depth in ‘hertzfit’, this implies that cells are stable on Sample 2. The reason might be related to

focal adhesion. Ghibaudo et. al., [89] showed that fibroblast shows strong dependency of

adhesion on spacing between the pillars. But in this study, the spacing between the pillars are

the same. But yet, Kuo et al., [90] showed that focal adhesion is dependent on size of the pillars


and the cell-line. Authors worked with different cell lines such as CHO, MDCK and C2C12, and

observed their interaction with different size of the pillars, 200nm and 700nm. Authors

discovered that focal adhesions decreased on small pillars and mentioned that Chien et al.

showed that when cells have small focal adhesion they exert stronger force because small focal

complex become matured to large focal adhesion and thus exert contractile force on the

substrate. This exerted force on the substrate cause the pillars to bend. Moreover, Yim et. al.,

[91] showed s similar result. Therefore, it can be said that because of the small focal adhesion

form on small size of the pillars, Sample 2 possesses large force hence, highest stiffness. Biggs et.

al. [92] mentioned that when height of the nano-feature is small the focal adhesion increase. As

it pointed out before (2.2 section), cells form focal adhesion through integrin and in 3T3

fibroblast there are three different types of integrin: α5β1, α5β3, clone of α5β1/α5β3. They recruit

focal adhesion molecules and form strong focal adhesion with fibronectin which is a component

of ECM, in Sample 2 compared to other Samples.

To find out the statistical significance of these results, p value was determined pairwise. The null

hypothesis, h values are 1 and 0. When h=0, the test failed to reject the null hypothesis at the

5% significant level and vice versa when h=1. Here are the P and h value for 0.4-0.6 µm height

are:

S1-S2: P = 4.81*10-5 (h=1), S1-S3b: P = 0.009 (h=1), S2-S3b: P = 0.125 (h=0);

From these value it is seen that only the S2-S3b null hypothesis is significant.

On the other hand for 1.9- 2.1 µm height the P and h values are:

S1-S2: P = 0.023 (h=0), S2-S3b: P = 0.002 (h=1) and S1-S3b: P = 0.055 (h=0);

From these it is observed that S2-S3b pair statistic is not significant but other two, S1-S2, S1-S3b

are significant.

In 0.9-1.1 and 1.4-1.6 µm height none of them are significant.

S1-S2: P = 1.02*10-4 (h=1), S1-S3b: P = 0.02 (h=1), S2-S3b: P = 0.015 (h=1); (0.9-1.1µm);

S1-S2: P = 0.007 (h=1), S1- S3b: P = 0.014 (h=1), S2-S3b: P = 0.003 (h=1); (1.4-1.6µm).

Individual measuerements of Young’s modulus over height of the cells (left) and the substrates

(right) for 3 samples represented in figure 4-24. All of them show decrease of elasticity with

increasing height for cells however, for the substrates, it is steady.


Figure 4-24: 3D plot of Young’s modulus over height of 3 samples, colour scale bar shows how frequent the combination of young’s modulus and height was measured, left figures are for the cell and right figures are for the substrate.

The most frequent elastic modulus for cells were for Sample 1 : 5-3kPa in 0.5- 0.9 µm height, for

Sample 2: 10-7kPa in 0-1 µm height and for Sample 3b: 10-6kPa between 0.6 and 0.8 µm height.

It can be seen that lowest part of the cell was the stiffest part and it corresponded to edge of the

cell. From figure 4-24 it is noticed that cell heights were different for most frequent elasticity,

which might be due to glueing the substrates onto the petri dishes.

Ning et. el., [93] did similar work with the specification: distance between the pillars was 700nm,

diameter of pillars was 200nm and height was 300nm. The stiffness for 3T3 fibroblast at an

indentation depth of 300nm was found on flat surface was 2.4kPa and on nano-pillars was

1.5kPa. The stiffness was found higher in Nano-channels (distance 555-diameter 150- height

140nm) is 2.25kPa. In contrast, in this study the cells were stiffer on the pillars than on the flat

surface, Table 4-2. Therefore, it can be said that when the substrates have more groove the

Sample 3b (5 cells measurements)

Sample 2 (30 cells measurements)

Sample 1 (10 cells measurements)


elasticity of the cell will increase. Additionally, a study [16] stated that the elasticity of 3T3

fibroblast is 140±30 dyne/cm2 (14±3Pa) which is much smaller than this study.

Table 4-2: Elastic modulus with different Samples and Spring Constant

Sample type Calibrated Spring

constant (k) mN/m

Elasticity of

cell (kPa)

Elasticity of

substrate (MPa)

Sample 1 (130-52-106) 276.1 3.7 (10 cells) 1

Sample 2 (130-34-75) 124.6 10 (30 cells) 0.5

Sample 3a (130-52-75) 83.94 1 (6 cells) 0.13

Sample 3b (130-52-75) 276.1 4.3 (5 cells) 1

For Hertz model, it assumed that cells are homogeneous and this model tells about the static

Young’s modulus but not the dynamic young’s modulus. Furthermore, cells are heterogeneous

so the evaluation of Young’s modulus using Hertz model may give an error. Also, cells have

viscoelastic property but the Hertz model neglects this.

It is mentioned before that AFM measurements rely on the spring constant (k) therefore, it is

very important to do the calibration of the cantilever carefully otherwise, the results will be

incorrect. Some of experimental error might have interrupted the calibration, thus in current

study there are different values for the spring constant. The probable reasons are:

Cantilever can be contaminated, thus increasing spring constant (k) value;

The liquid medium might have different density therefore k value will change;

The laser focused on the cantilever might be in different position that’s changes the

sensitivity;

For thermal noise analysis, temperature is one of the parameters, which can also change

k value etc.

CELL-SURFACE INTERACTION

Figure 4-25 shows the AFM picture of Sample 1, built by JPK software. With AFM It is possible to

see topography but not the pillars because the samples cannot be tilted. Moreover, the principle

of AFM is different from electron microscopy. The principle is already explained in experimental

part so it will not be repeated here. Therefore, to see cell-surface interaction, HRSEM (Hitachi

5200) was used. The samples for HRSEM were prepared by the Electron Microscopy

Department, assisted with Professor Paul Walther.


Figure 4-25: AFM picture (topography) of Sample 1 after background corrections, scale 5µm, colour scale bar shows the height measured

In section 2.2 it is stated that fibroblast moves smoothly and makes elongated triangle,

lamellipodia form on one side and extend forward. When lamellipodia moves it makes focal

adhesion to the substrate but it also detached when it reaches its proximal position. Apparently,

in figure 4-25, it is seen that cells are attached to the surface and spread very well regardless of

the height or the diameter of the pillar. Cells are very flat on the surface. Generally, when cells

are loosely attached to the surface they look spherical [94]. In this case cells were not spherical


so cells are firmly attached. It was also observed that when cell moves it only attaches to the top

of the pillars as shown in black arrows, figure 4-27.

Figure 4-26:HRSEM pictures at 5kV, Cells spread over the surface of all types of samples; a) Elongated triangular shape with formation of microvili, scale 20µm, b) without microvili, scale 30µm c) the formation of lamellipodia and filopodia (yellow arrows), scale 5µm

a) b)

c)


Figure 4-27: HRSEM picture taken at 5kV, 30 degree tilted, Cell moves attaching the top of the pillars in all samples (black arrows) a) Sample 2 scale 100nm, b) Sample 1, scale 1µm and c) Sample 3b 1µm.

a) a) b)

c)

Chapter 5: Conclusion Page 57 of 63

Chapter 5. CONCLUSION

In this study the elasticity of 3T3 fibroblast was investigated on Silicon nanostructures.

Nanostructure was prepared by both conventional methods: bottom up and top down

techniques. Dip coating process, which is a bottom up technique, used to make thin film with

constant withdrawal speed using block copolymer micellar gold solution and subsequent H2

plasma gives the deposition and interparticle distance of 130nm of the Au NPs. From here

hexagonally ordered gold nanoparticles were created with 9nm average diameter of Au NPs.

From controlled photochemical growth it was able to make Au NPs bigger in size to 17nm and

30nm with exposure time 1.5 and 3.5 minutes respectively. After conducting RIE which is top

down technique, the cylindrical like and hexagonally ordered nanopillars were produced with

different heights (75nm and 106nm) and diameters (28nm and 49nm) with aspect ratio, width

over height 0.52. This study showed that stiffness of cells are higher when diameter and height

of the pillars are small. The most frequent elastic modulus belongs to actin filaments. It was

observed that with increasing thickness of the cell the Young’s modulus decreased and different

heights of the pillars do not have significant influence on elasticity for the substrate and cells. It

is discovered that the cell was spread over the surface regardless of pillar heights or diameters.

Depending on the shape of the tip and calibration data of cantilever, the measurement will be

changed so it is necessary to use one spring constant to evaluate all data. Furthermore,

depending upon the model applied for the measurement the results will vary. Hertz model was

used to evaluate the data and this model is based on some assumptions. An evaluation can be

carried out with Finite Element Analysis (FEA) and results can be compared. Additionally, it can

be compared with cancerous fibroblast cell or even treated cells with inhibitor. Finally, since

living cells are interacting with nano-sized pillars and cells exert focal adhesion, there will be

pulling of the pillars therefore, the pillars might tilted and deformation of the pillars can be

determined.

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Chapter 6. REFERENCES

[1] W. Callister and D. Rethwisch, Materials science and engineering: an introduction, 7th ed. York, PA: John Wiley & Sons, 2007.

[2] V. K. Varadan, A. S. Pillai, D. Mukherji, M. Dwivedi, and L. Chen, Nanoscience and Nanotechnology in Engineering, 1st ed. World Scientific, 2010.

[3] A. Sunny and S. Thomas, “Nanoparticles,” in Recent Advances in Polymer Nanocomposites, CRC Press, 2010, pp. 1–18.

[4] G. L. Hornyak, H. F. Tibbals, J. Dutta, and J. J. Moore, Introduction to Nanoscience and Nanotechnology. 2009.

[5] M. Köhler and W. Fritzsche, “Microtechnological Foundations,” in Nanotechnology: An Introduction to Nanostructuring Techniques, 2nd ed., Wiley-VCH, 2007, pp. 33–85.

[6] F. J. S. and J. E. L. Buddy D. Ratner, Allan S. Hoffman, B. D. Ratner, A. S. Hoffman, F. J. Schoen, and J. E. Lemons, Biomaterials Science, 3rd Edition An Introduction to Materials in Medicine, no. 1. 2001.

[7] J. Park and R. S. Lakes, Biomaterials: An Introduction, 3rd ed. New York: Springer New York, 2007.

[8] J. J. Ramsden, “Chapter 1 - What is Nanotechnology?,” in Nanotechnology: An Introduction. A volume in Micro and Nano Technologies, Oxford: William Andrew Publishing, 2011, pp. 1–14.

[9] A. Colas and J. Curtis, “Silicone biomaterials: history and chemistry,” Biomater. Sci. an Introd. to Mater. Med., vol. 2, pp. 80–85, 2004.

[10] L. Sirghi, “Atomic Force Microscopy indentation of living cells,” Microsc. Sci. Technol. Appl. Educ., vol. 4, pp. 433–440, 2010.

[11] M. J. Dalby, D. Giannaras, M. O. Riehle, N. Gadegaard, S. Affrossman, and A. S. G. Curtis, “Rapid fibroblast adhesion to 27 nm high polymer demixed nano-topography,” Biomaterials, vol. 25, no. 1, pp. 77–83, 2004.

[12] L. Yang, H. Liu, and Y. Lin, “Biomaterial nanotopography-mediated cell responses: experiment and modeling,” Int. J. Smart Nano Mater., vol. 2014, no. January, 2014.

[13] Y. Wang and D. E. Discher, Eds., Cell Mechanics. Academic Press, 2007.

[14] A. note JKP, “Determining the elastic modulus of biological samples using atomic force microscopy,” pp. 1–9.

[15] M. Radmacher, M. Fritz, C. M. Kacher, J. P. Cleveland, and P. K. Hansma, “Measuring the viscoelastic properties of human platelets with the atomic force microscope.,” Biophys. J., vol. 70, no. 1, pp. 556–67, Jan. 1996.

[16] A. Tondon, C. Haase, and R. Kaunas, “Mechanical Stretch Assays in Cell Culture Systems,” in Handbook of Imaging in Biological Mechanics, C. P. N. Guy and M. Genin, Eds. CRC Press, 2014, pp. 313–322.

[17] B. Alberts, D. Bray, J. Lewis, M. Raff, K. Roberts, and J. D. Watson, Molecular Biology of the Cell, 2nd ed. New York: Garland Publishing, Inc, 1989.

[18] B. Alberts, Molecular biology of the cell, 6th ed. New York: Garland Publishing, Inc, 2015.

[19] B. Alberts, A. Johnson, and J. Lewis, “The Extracellular Matrix of Animals,” in Molecular Biology of the Cell, vol. i, 2002.

[20] C. Frantz, K. M. Stewart, and V. M. Weaver, “The extracellular matrix at a glance.,” J. Cell Sci., vol.

of 63

123, pp. 4195–4200, 2010.

[21] B. Alberts, A. Johnson, J. Lewis, M. Raff, K. Roberts, and P. Walter, “Fibroblasts and their transformations: the connective-tissue cell family,” in Molecular Biology of the Cell, 4th ed., New York: Garland Science, 2002.

[22] G. M. Cooper, The Cell: A Molecular Approach, 2nd ed. Sunderland, MA: Sinauer Associates, 2000.

[23] D. H. Boal, “Polymers,” in Mechanics of the Cell, 1st, Ed. Cambridge: Cambridge University Press, 2002, pp. 23–58.

[24] D. A. Fletcher and R. D. Mullins, “Cell mechanics and the cytoskeleton,” Nature, vol. 463, no. 7280, pp. 485–492, Jan. 2010.

[25] M. L. Rodriguez, P. J. McGarry, and N. J. Sniadecki, “Review on Cell Mechanics: Experimental and Modeling Approaches,” Appl. Mech. Rev., vol. 65, no. 6, p. 60801, Oct. 2013.

[26] B. Alberts, A. Johnson, J. Lewis, M. Raff, K. Roberts, and P. Walter, “Integrins,” in Molecular Biology of the Cell, 4th ed., New York: Garland Science, 2002, p. 1616 pages.

[27] D. Bray, Cell Movements: From Molecules to Motility, 2nd ed. New York: Garland Publishing, Inc, 2001.

[28] G. M. Cooper, “Cell Crawling,” in The Cell: A Molecular Approach, 2nd ed., Sunderland, MA: Sinauer Associates, 2000.

[29] M. M. Bouziane, S. Benbarek, E. M. H. Tabeti, B. A. B. Bouiadjra, N. Benseddiq, B. Serier, and A. Albedah, “Design and Computation of Modern Engineering Materials,” A. Öchsner and H. Altenbach, Eds. Cham: Springer International Publishing, 2014, pp. 73–87.

[30] M. Hervy, “Modulation of Cell Structure and Function in Response to Substrate Stiffness and External Forces,” J. Adhes. Sci. Technol., vol. 24, no. 5, pp. 963–973, Jan. 2010.

[31] S. Timoshenko and J. N. Goodier, “Theory of Elasticity,” Journal of Elasticity, vol. 49. pp. 427–143, 1986.

[32] A. Vinckier and G. Semenza, “Measuring elasticity of biological materials by atomic force microscopy,” FEBS Lett., vol. 430, no. 1–2, pp. 12–16, 1998.

[33] V. L. Popov, “Kontaktmechanik und Reibung,” J. Chem. Inf. Model., vol. 53, pp. 1689–1699, 2009.

[34] H. Ladjal, J. Hanus, A. Pillarisetti, C. Keefer, A. Ferreira, and J. P. Desai, “Atomic Force Microscopy-Based Single-Cell Indentation : Experimentation and Finite Element Simulation,” pp. 1326–1332, 2009.

[35] J. A. Last, P. Russell, P. F. Nealey, and C. J. Murphy, “The applications of atomic force microscopy to vision science,” Investig. Ophthalmol. Vis. Sci., vol. 51, no. 12, pp. 6083–6094, 2010.

[36] M. Radmacher, “Atomic Force Microscopy in Cell Biology,” Elsevier, p. 82.

[37] I. Clean Air Technology, “What is a Cleanroom?” [Online]. Available: http://www.cleanairtechnology.com/cleanroom-classifications-class.php. [Accessed: 19-May-2016].

[38] Simplex Technical Staff, “What to Know When Considering a Cleanroom,” Fontana, CA, 2015.

[39] U.S. Federal Standard, “Cleanroom specification,” 2013. [Online]. Available: http://www.cmgreenbuilding.com/clean-rooms/. [Accessed: 23-Apr-2016].

[40] S. L. Hellstrom, “Basic Models of Spin Coating,” Coursework Phys. 210, Stanford Univ., vol. 2007, pp. 1–6, 2007.

[41] A. G. Emslie, F. T. Bonner, and L. G. Peck, “Flow of a Viscous Liquid on a Rotating Disk,” J. Appl. Phys., vol. 29, no. 5, p. 858, 1958.

of 63

[42] K. Norrman, A. Ghanbari-Siahkali, and N. B. Larsen, “6 Studies of spin-coated polymer films,” Annu. Reports Sect. “C” (Physical Chem., vol. 101, p. 174, Oct. 2005.

[43] N. P. Pham, E.Boellard, P. M. Sarro, and J. N. Burghartz, “Spin, Spray coating and Electrodeposition of photoresist for MEMS structures – A comparison,” Eurosensors, pp. 81–86, 2002.

[44] “Spin Coating,” Basic Laboratory Materials: Science and&Engineering, 2015. [Online]. Available: http://www.tf.uni-kiel.de/servicezentrum/en/studium/praktika/basic. [Accessed: 15-Feb-2016].

[45] C. J. Brinker, “Dip Coating,” in Chemical Solution Deposition of Functional Oxide Thin Films SE - 10, T. Schneller, R. Waser, M. Kosec, and D. Payne, Eds. Springer Vienna, 2013, pp. 233–261.

[46] J. Puetz and M. A. Aegerter, “Sol-Gel Technologies for Glass Producers and Users,” M. A. Aegerter and M. Mennig, Eds. Boston, MA, MA: Springer US, 2004, pp. 37–48.

[47] G. Kästle, H.-G. Boyen, F. Weigl, P. Ziemann, S. Riethmüller, C. H. Hartmann, J. P. Spatz, M. Möller, M. G. Garnier, and P. Oelhafen, “The Self-organization of Metal Loaded Micelles - An Approach to Prepare Ordered Arrays of Metallic Nanoislands,” Phase Transitions, vol. 76, no. 4–5, pp. 307–313, Jan. 2003.

[48] H. Ji, G. Sakellariou, R. C. Advincula, G. D. Smith, S. M. Kilbey, M. D. Dadmun, and J. W. Mays, “Synthesis and characterization of well-defined [polystyrene-b-poly(2-vinylpyridine)]n star-block copolymers with poly(2-vinylpyridine) corona blocks,” J. Polym. Sci. Part A Polym. Chem., vol. 45, no. 17, pp. 3949–3955, Sep. 2007.

[49] B. Ö. and W. H. and A. P. and P. Ziemann, “Arrays of quasi-hexagonally ordered silica nanopillars with independently controlled areal density, diameter and height gradients,” Nanotechnology, vol. 26, no. 11, p. 115301, 2015.

[50] G. Kästle, H.-G. Boyen, F. Weigl, G. Lengl, T. Herzog, P. Ziemann, S. Riethmüller, O. Mayer, C. Hartmann, J. P. Spatz, M. Möller, M. Ozawa, F. Banhart, M. G. Garnier, and P. Oelhafen, “Micellar Nanoreactors—Preparation and Characterization of Hexagonally Ordered Arrays of Metallic Nanodots,” Adv. Funct. Mater., vol. 13, no. 11, pp. 853–861, Nov. 2003.

[51] J. Chai, D. Wang, X. Fan, and J. M. Buriak, “Assembly of aligned linear metallic patterns on silicon,” Nat Nano, vol. 2, no. 8, pp. 500–506, Aug. 2007.

[52] G. Bruno, M. Losurdo, and P. Capezzuto, “On the Use of H2 Plasma for the Cleaning and Passivation of InP Substrates,” J. Phys. IV Fr., vol. 05, no. C5, pp. C5–663–C5–670, Jun. 1995.

[53] K. S. Choi and C. M. Lee, “Removal efficiency of organic contaminants using ECR H 2 plasma and ECR O 2 plasma,” 2003.

[54] T. H. and A. S. and P. O. and A. P. and P. Z. and L. M. Eng, “Controlled photochemical particle growth in two-dimensional ordered metal nanoparticle arrays,” Nanotechnology, vol. 21, no. 14, p. 145309, 2010.

[55] H. Hsu and E. Engineering, “Photochemical Synthesis of Gold Nanoparticles with Interesting Shapes Abstract : Introduction :,” Reactions, pp. 68–69, 2004.

[56] K. Mallick, Z. L. Wang, and T. Pal, “Seed-mediated successive growth of gold particles accomplished by UV irradiation: a photochemical approach for size-controlled synthesis,” J. Photochem. Photobiol. A Chem., vol. 140, no. 1, pp. 75–80, 2001.

[57] “Suss MJB-3 Operator’s Manual,” Garching, 2004.

[58] B. Özdemir, “Systematically Varied Arrangements and Geometries of SiO2 Nanopillars serving as Static and Dynamic Platforms for studying Cell Responses,” Universität Ulm, 2015.

[59] A. Nayak, M. S. Islam, and V. J. Logeeswaran, “Wet Etching,” in Encyclopedia of Nanotechnology SE - 431, B. Bhushan, Ed. Springer Netherlands, 2012, pp. 2829–2830.

[60] H. Jansen, H. Gardeniers, M. de Boer, M. Elwenspoek, and J. Fluitman, “A survey on the reactive

of 63

ion etching of silicon in microtechnology,” J. micromechanics microengineering, vol. 6, no. 1, p. 14, 1996.

[61] S. Franssila, “Etching,” in Introduction to Microfabrication, John Wiley & Sons, Ltd, 2010, pp. 127–141.

[62] C. J. Mogab, A. C. Adams, and D. L. Flamm, “Plasma etching of Si and SiO2 - The effect of oxygen additions to CF4 plasmas,” J. Appl. Phys., vol. 49, no. 7, pp. 3796–3803, 1978.

[63] M.Elwenspoek & H.V. Jansen, Silicon Micromatching. Cambridge University Press, 1998.

[64] C. Cantafio, “How the SEM Works,” Scanning Electron Microscope Usage and Analysis of Metal Fatigue in Mead Silver Paperclips, 1998. [Online]. Available: http://antoine.frostburg.edu/engin/sem/workings.html. [Accessed: 23-Apr-2016].

[65] JEOL Ltd., “Scanning Electron Microscope A To Z,” pp. 1–32.

[66] W. C. Nixon, “The General Principles of Scanning Electron Microscopy,” Jstor, vol. 261, no. 837, pp. 45–50, 1971.

[67] C. Cantafio, “How the SEM Works,” Scanning Electron Microscope Usage and Analysis of Metal Fatigue in Mead Silver Paperclips, 1998. .

[68] I. Sokolov, “Atomic force microscopy in cancer cell research,” Cancer Nanotechnol., vol. 1, pp. 1–17, 2007.

[69] F. L. Leite, L. H. C. Mattoso, O. N. O. Jr, and P. S. P. H. Jr, “The Atomic Force Spectroscopy as a Tool to Investigate Surface Forces : Basic Principles and Applications,” Mod. Res. Educ. Top. Microsc., pp. 22–24, 2007.

[70] R. Howland and L. Benatar, “A Practical Guide To Scanning Probe Microscopy,” p. 87, 1996.

[71] JPKInstruments, “A practical guide to AFM force spectroscopy and data analysis,” Tech Sheet, pp. 1–8.

[72] W. Han and M. Serry, “Force Spectroscopy with the Atomic Force Microscope,” pp. 1–8, 2008.

[73] A. F. Microscopy and A. Jpk, “QITM mode - Quantitative Imaging with the NanoWizard ® 3 AFM,” pp. 1–7.

[74] G. Berteloot, C.-T. Pham, a. Daerr, F. Lequeux, and L. Limat, “Evaporation-induced flow near a contact line: Consequences on coating and contact angle,” EPL (Europhysics Lett., vol. 83, no. 1, p. 14003, 2008.

[75] J. Bansmann, S. Kielbassa, H. Hoster, F. Weigl, H. G. Boyen, U. Wiedwald, P. Ziemann, and R. J. Behm, “Controlling the Interparticle Spacing of Au−Salt Loaded Micelles and Au Nanoparticles on Flat Surfaces,” Langmuir, vol. 23, no. 20, pp. 10150–10155, Sep. 2007.

[76] T. K. Sau, A. Pal, N. R. Jana, Z. L. Wang, and T. Pal, “Size controlled synthesis of gold nanoparticles using photochemically prepared seed particles,” J. Nanoparticle Res., vol. 3, no. 4, pp. 257–261, 2001.

[77] B. Özdemir, A. Seidenstücker, A. Plettl, and P. Ziemann, “Cyclic photochemical re-growth of gold nanoparticles: Overcoming the mask-erosion limit during reactive ion etching on the nanoscale,” Beilstein J. Nanotechnol., vol. 4, no. 1, pp. 886–894, 2013.

[78] S. Brieger, O. Dubbers, S. Fricker, A. Manzke, C. Pfahler, A. Plettl, and P. Ziemann, “An approach for the fabrication of hexagonally ordered arrays of cylindrical nanoholes in crystalline and amorphous silicon based on the self-organization of polymer micelles,” Nanotechnology, vol. 17, no. 19, pp. 4991–4994, 2006.

[79] N. Cheung, “Reactive Ion Etching (RIE): Lecture 15,” 2010.

[80] O. V. Balachova, M. A. R. Alves, J. W. Swart, E. S. Braga, and L. Cescato, “CF4 plasma etching of

of 63

materials used in microelectronics manufacturing,” Microelectronics J., vol. 31, no. 3, pp. 213–215, 2000.

[81] B. Christian, “Bestimmung der Steifigkeit von Zellen aus Kraft-Distanz-Kurven,” Ulm University.

[82] H. Haga, S. Sasaki, K. Kawabata, E. Ito, T. Ushiki, and T. Sambongi, “Elasticity mapping of living "Fibroblasts by AFM and immunofluorescence observation of the cytoskeleton,” Ultramicroscopy, vol. 82, pp. 253–258, 2000.

[83] S. Y. Tee, J. Fu, C. S. Chen, and P. A. Janmey, “Cell shape and substrate rigidity both regulate cell stiffness,” Biophys. J., vol. 100, no. 5, pp. L25–L27, 2011.

[84] T. G. Kuznetsova, M. N. Starodubtseva, N. I. Yegorenkov, S. A. Chizhik, and R. I. Zhdanov, “Atomic force microscopy probing of cell elasticity,” Micron, vol. 38, no. 8, pp. 824–833, 2007.

[85] B. Codan, V. Martinelli, L. Mestroni, and O. Sbaizero, “Atomic force microscopy of 3T3 and SW-13 cell lines: An investigation of cell elasticity changes due to fixation,” Mater. Sci. Eng. C, vol. 33, no. 6, pp. 3303–3308, 2013.

[86] J. Solon, I. Levental, K. Sengupta, P. C. Georges, and P. A. Janmey, “Fibroblast Adaptation and Stiffness Matching to Soft Elastic Substrates,” Biophys. J., vol. 93, no. 12, pp. 4453–4461, 2007.

[87] S. Park, D. Koch, R. Cardenas, J. Käs, and C. K. Shih, “Cell motility and local viscoelasticity of fibroblasts.,” Biophys. J., vol. 89, no. 6, pp. 4330–42, 2005.

[88] C. Rotsch, K. E. N. Jacobsont, M. R. A. I. Macher, T. J. Meyer, N. Carolina, and C. I. Hill, “Dimensional and mechanical dyni tmics of active and stable edges in motile fibroblasts investigated I by using atomic force microscopy,” vol. 96, no. February, pp. 921–926, 1999.

[89] M. Ghibaudo, L. Trichet, J. Le Digabel, A. Richert, P. Hersen, and B. Ladoux, “Substrate topography induces a crossover from 2D to 3D behavior in fibroblast migration,” Biophys. J., vol. 97, no. 1, pp. 357–368, 2009.

[90] C. Kuo, D. Chueh, and P. Chen, “Investigation of size–dependent cell adhesion on nanostructured interfaces,” J. Nanobiotechnology, vol. 12, p. 54, 2014.

[91] E. K. F. Yim and K. W. Leong, “Significance of synthetic nanostructures in dictating cellular response,” Nanomedicine Nanotechnology, Biol. Med., vol. 1, no. 1, pp. 10–21, 2005.

[92] M. J. P. Biggs, R. G. Richards, and M. J. Dalby, “Nanotopographical modification: A regulator of cellular function through focal adhesions,” Nanomedicine Nanotechnology, Biol. Med., vol. 6, no. 5, pp. 619–633, 2010.

[93] D. Ning, B. Duong, G. Thomas, Y. Qiao, L. Ma, Q. Wen, and M. Su, “Mechanical and morphological analysis of cancer cells on nanostructured substrates,” Langmuir, p. acs.langmuir.5b04469, 2016.

[94] M. E. Dokukin, N. V. Guz, and I. Sokolov, “Quantitative study of the elastic modulus of loosely attached cells in AFM indentation experiments,” Biophys. J., vol. 104, no. 10, pp. 2123–2131, 2013.

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