Near-field infrared nanoscopy applied to laterally structured self

Transcription

Near-field infrared nanoscopy
applied to
laterally structured
self-assembled monolayers
Dissertation
submitted for the degree of
Dr. rer. nat. (Doctor rerum Naturalium)
in the faculty of chemistry and biochemistry
at the Ruhr-University Bochum
Germany
Dipl.-Biochem. Ilona Kopf
Department of Physical Chemistry II
Bochum, 2008
The work was carried out between October 2004 and November 2008 at the department
for Physical Chemistry II under the supervision of Prof. Dr. M. Havenith.
1st Examiner: Prof. Dr. M. Havenith
2nd Examiner: Prof. Dr. Ch. Wöll
Thesis Committee Head: Prof. Dr. R. Heumann
Date of Defense: 12/01/2009
Declaration
I hereby declare that the dissertation entitled ”Near-field infrared nanoscopy applied to
laterally structured self-assembled monolayers” is my original work and has been written
with no other sources and aids than quoted, and has not been submitted elsewhere for
an examination, as thesis or for evaluation in a similar context.
Ilona Kopf
für meine Mutter und meine Schwester
Holzhacken ist deswegen so beliebt,
weil man bei dieser Tätigkeit
den Erfolg sofort sieht
Albert Einstein
Wir müssen unbedingt Raum für Zweifel lassen,
sonst gibt es keinen Fortschritt, kein Dazulernen.
Man kann nichts Neues herausfinden,
wenn man nicht vorher eine Frage stellt.
Und um zu fragen, bedarf es des Zweifels.
Richard P. Feynman
Contents
1. Introduction and outline of thesis
1
2. Theory of self-assembled monolayers (SAMs)
2.1. Molecular self-assembly . . . . . . . . . . .
2.2. Brief historical introduction to SAMs . . .
2.3. Basic theory of thiolate-gold SAMs . . . .
2.4. DNA SAMs - a special case . . . . . . . .
on
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3. Theory of lithographic techniques to laterally structure SAMs
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3.1. Microcontact printing (µCP) . . . . . . . . . . . . . . . . . . . . . . . . . 22
3.2. Scanning probe lithography (SPL) . . . . . . . . . . . . . . . . . . . . . . 26
4. Theory of scanning probe microscopy (SPM)
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4.1. Atomic force microscopy (AFM) . . . . . . . . . . . . . . . . . . . . . . . 32
4.2. Chemical force microscopy (CFM) . . . . . . . . . . . . . . . . . . . . . . 37
5. Theory of infrared (IR) spectroscopy
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5.1. Infrared spectroscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 40
5.2. Fourier transform infrared (FTIR) spectroscopy . . . . . . . . . . . . . . 44
5.3. IR microscopy or microspectroscopy . . . . . . . . . . . . . . . . . . . . . 48
6. Theory of scanning near-field optical microscopy (SNOM)
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6.1. Historical development of SNOM . . . . . . . . . . . . . . . . . . . . . . 53
6.2. SNOM configurations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55
6.3. Theory of s-SNOM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 60
7. Preparation and characterization of
7.1. Materials and Methods . . . . .
7.2. Results and Discussion . . . . .
7.3. Conclusion . . . . . . . . . . . .
gold
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substrates
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8. Stamp fabrication and microcontact printing (µCP) of ODT
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8.1. Materials and Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . 84
8.2. Results and Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86
8.3. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 89
I
Contents
9. Experimental set-up of the scattering scanning near-field infrared microscope
(s-SNIM)
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9.1. Scattering scanning near-field microscope . . . . . . . . . . . . . . . . . . 92
9.2. Carbon monoxide (CO) laser . . . . . . . . . . . . . . . . . . . . . . . . . 97
10.Characterization of microstructured monolayers of biotinylated alkylthiolates
by s-SNIM
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10.1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 104
10.3. Results and Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . 106
11.Detection of hybridization in DNA monolayers by IRRAS, AFM and s-SNIM117
11.1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 118
11.3. Characterization of DNA SAMs by IRRAS . . . . . . . . . . . . . . . . . 128
11.4. Characterization of nanografted DNA and DNA SAMs using AFM . . . . 139
11.5. Characterization of DNA nanostructures using s-SNIM . . . . . . . . . . 152
11.6. Summarizing conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . 157
12.Infrared dyes in s-SNIM
12.1. Introduction . . . . . .
12.2. Materials and Methods
12.3. Results and Discussion
12.4. Conclusion . . . . . . .
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13.Bibliography
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14.Acknowledgements
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A. Table CO laser lines
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B. Typical vibrational bands of biomolecules
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C. Purchasing of synthetic polynucleotides
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D. Symbols and Abbreviations
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E. Curriculum Vitae and Publications
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II
List of Figures
2.1. Schematic diagram of a SAM . . . . . . . . . . . .
2.2. Different methods for monolayer deposition on solid
2.3. Schematic diagram of an alkylthiolate SAM . . . .
2.4. Orientation of a long-chain alkanethiolate on gold .
2.5. SAM growth in solution . . . . . . . . . . . . . . .
2.6. Schematic structure of a DNA nucleotide . . . . . .
2.7. Schematic structure of a DNA double strand . . . .
2.8. Schematic structure of a DNA helix . . . . . . . . .
2.9. Scheme of a secondary structured hairpin DNA . .
2.10. Chemical structure of 5’-thiolated DNA . . . . . . .
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3.1.
3.2.
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4.1. AFM operation modes . . . . . . . . . . . . . . . . . . . . . . . . . . . .
4.2. AFM laser beam deflection system and AFM set-up . . . . . . . . . . . .
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5.1.
5.2.
5.3.
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6.1. Synge’s concept on near-field microscopy . . . . . . . . . . . . . . . . . .
6.2. Aperture and scattering SNOM . . . . . . . . . . . . . . . . . . . . . . .
6.3. Effect of SNOM probe’s cone angles on light propagation and topographic
contrast . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.4. Aperture SNOM operation modes . . . . . . . . . . . . . . . . . . . . . .
6.5. s-SNOM operation modes . . . . . . . . . . . . . . . . . . . . . . . . . .
6.6. s-SNOM theory: the tip . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.7. s-SNOM theory: tip-sample interaction . . . . . . . . . . . . . . . . . . .
6.8. s-SNOM theory: Orientation of the external electric field . . . . . . . . .
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Polymerization reaction of PDMS . . . . . . . . . .
PDMS stamp fabrication and microcontact printing
Further treatment of laterally structured SAMs . .
Different types of stamp deformations . . . . . . . .
Nanografting process . . . . . . . . . . . . . . . . .
Harmonic and anharmonic oscillator . . .
Typical molecular vibration modes . . . .
Dispersive and FTIR spectrometer . . . .
Signal processing in a FTIR spectrometer .
Surface selection rule . . . . . . . . . . . .
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III
List of Figures
6.9. Near-field theory: Background signal . . . . . . . . . . . . . . . . . . . .
6.10. Interferometric near-field detection schemes . . . . . . . . . . . . . . . .
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7.1. Preparation of template stripped gold . . . . . . . . . . . . . . . . . . . .
7.2. Topographs of sputter deposited gold and TSG . . . . . . . . . . . . . .
7.3. Efficiency of piranha cleaning (IRRAS spectra) . . . . . . . . . . . . . . .
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8.1. Scheme and dimensions of AFM calibration grids . . . . . . . . . . . . .
8.2. Stamp casting and subsequently cutting of excess PDMS . . . . . . . . .
8.3. Light and AFM topograph of a stamp fabricated from a TGZ11 calibration grid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
8.4. Dynamic mode AFM topographs of printed ODT patterns from self-made
stamps . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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9.1.
9.2.
9.3.
9.4.
IV
s-SNIM set-up and beam path
s-SNIM detection unit . . . .
CO-laser set-up . . . . . . . .
CO-laser emission spectra . .
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10.1. Molecular structures of BAT and ODT . . . . . . . . . . . . . . . .
10.2. SEM photomicrograph of PDMS stamp . . . . . . . . . . . . . . . .
10.3. SEM photomicrograph of patterned BAT-ODT SAM . . . . . . . .
10.4. FTIR spectra of BAT . . . . . . . . . . . . . . . . . . . . . . . . . .
10.5. IRRAS spectra of BAT and ODT . . . . . . . . . . . . . . . . . . .
10.6. Topograph and near-field image of a BAT-ODT SAM at 1711 cm−1
10.7. Near-field contrast of BAT as function of wavenumber . . . . . . . .
10.8. Height artifacts in s-SNIM . . . . . . . . . . . . . . . . . . . . . . .
10.9. Lateral resolution of s-SNIM . . . . . . . . . . . . . . . . . . . . . .
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11.1. Secondary structures of probe and target ssDNA . . . . . . . . . . . . . .
11.2. Scheme of the nanografting process . . . . . . . . . . . . . . . . . . . . .
11.3. Scheme of the nanografting process . . . . . . . . . . . . . . . . . . . . .
11.4. Determination of the AFM sensitivity . . . . . . . . . . . . . . . . . . . .
11.5. Calculated IR spectra for DNA bases and base pairs . . . . . . . . . . . .
11.6. Calculated IR spectra for the employed DNA sequence . . . . . . . . . .
11.7. IRRAS spectra of MCH adsorbed from STE-buffer solution and ethanolic
solution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
11.8. Influence of MCH backfilling on IRRAS spectra of DNA SAMs . . . . . .
11.9. Comparison between IRRAS spectra of MCH, ssDNA and dsDNA SAMs
11.10.IRRAS spectra of DNA SAMs before and after hybridization . . . . . .
11.11.IRRAS spectrum of biotin labeled dsDNA . . . . . . . . . . . . . . . . .
11.12.IRRAS spectra of fresh and aged DNA SAMs . . . . . . . . . . . . . . .
11.13.Alteration of DNA SAM response as function of evacuation time . . . .
11.14.AFM height measurements of nanostructured DNA before and after hybridization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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List of Figures
11.15.Scheme on unconstrained assembly and SCSA of DNA . . . . . . . . . .
11.16.Theoretical predicted length of the employed DNA sequence . . . . . . .
11.17.AFM height measurements after different nanografting steps . . . . . . .
11.18.Compressibility of DNA after different nanografting steps . . . . . . . .
11.19.Kretchman-Raether configuration and SPR resonance curve . . . . . . .
11.20.SPR measurements on a low density DNA SAM . . . . . . . . . . . . . .
11.21.DNA melting curves and UV absorbance spectra . . . . . . . . . . . . .
11.22.Topographic images of nanografted DNA structures in air . . . . . . . .
11.23.s-SNIM measurements on nanografted DNA structures . . . . . . . . . .
11.24.s-SNIM contrast as function of probe density and hybridization efficiency
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12.1. Molecular structure of the cymantrene labeled peptide (CymPntCys) .
12.2. IRRAS spectra comparing peptide adsorption from ethanol and water .
12.3. Investigation of unspecific adsorption of CymPntCys on ODT . . . . .
12.4. AFM image of a laterally structured ODT-CymPntCys SAM . . . . . .
12.5. s-SNIM measurements of a laterally structures ODT-CymPntCys SAM
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V
List of Tables
2.1. Properties of principal components of molecules forming SAMs . . . . . .
2.2. Analytical techniques for SAM characterization . . . . . . . . . . . . . .
2.3. Comparison of geometries of the most common DNA forms . . . . . . . .
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5.1. Bond strength and vibrational frequency . . . . . . . . . . . . . . . . . .
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6.1. Resolution of different microscopy techniques . . . . . . . . . . . . . . . .
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7.1. Peak assignment for IRRAS spectra of ODT . . . . . . . . . . . . . . . .
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8.1. Parameter of AFM calibration grids used for stamp fabrication . . . . . .
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9.1. CO-laser gas composition . . . . . . . . . . . . . . . . . . . . . . . . . . .
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10.1. Peak assignment for IRRAS spectra of BAT and ODT . . . . . . . . . . 111
11.1. Oligonucleotide sequences . . . . . . . . . . . . . . . . . . .
11.2. Preparation of background SAMs for DNA nanografting . .
11.3. Peak assignment for calculated IR spectra of DNA bases and
11.4. Peak assignment for IRRAS spectra of MCH . . . . . . . . .
11.5. Peak assignment for IRRAS spectra of DNA . . . . . . . . .
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A.1. Laser transitions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 182
VII
1. Introduction and outline of thesis
Today some analytical techniques exists that enable high resolution sample characterization e.g. advanced fluorescence microscopy (stimulated emission depletion (STED), 4Pi
microscopy), electron microscopy and atomic force microscopy. But all of them require
either special sample preparation or extreme measurement conditions (e.g. UHV) or
they are not able to obtain chemical information.
Optical spectroscopy - in particular fingerprint infrared spectroscopy - is a powerful tool
for characterizing the chemical composition and structure of the species under investigation. Spatial resolved spectroscopy enables chemical mapping of samples. However,
in the traditional implementation of spatial resolved optical spectroscopy the resolution
is diffraction-limited to dimensions in the order of the wavelength. In case of infrared
spectroscopy the lateral resolution is especially worse due to the large wavelengths of
infrared radiation ranging from 800 nm to 1 mm. For many applications the achievable
resolution is absolutely insufficient e.g. studies of cellular differentiation, studies of subcellular compartments or applications in microelectronics.
A nice way to overcome these limitations is scattering near-field infrared microscopy a
technique breaking the diffraction limit by confining the light on the nanometer scale
using a suitable probe. This technique combines high resolution scanning probe microscopy with chemical sensitive, nanometer resolved infrared spectroscopy. Infrared
chemical mapping with nanometric resolution is desirable in many research fields of
material sciences and life sciences:
• semiconductors
• polymer research
• cellular and sub-cellular studies
• lipid phase separation (lipid rafts)
• membrane proteins
IR-SNOM performed in a time-resolved spectroscopic way can develop into a powerful
research tool for looking spectroscopically at ”hot spots” in biological system. For instance the activity of a membrane protein could be studied at the single molecule level
in situ. Thus a window to an entire new field of research is opened.
Beyond living systems also for further technological progress a nanoscopic infrared tool
is of great benefit. Future implantable biosensors are relying on biocompatible surfaces
at the macro- and nanometer scale which will need to be characterized in detail. SNIM
also seems to be ideally suited for characterizing nanopattered surfaces which are currently attracting a lot of interest due to their potential in improving biocompatibility of
1
Chapter 1. Introduction and outline of thesis
surfaces on various length scales.
The focus of this thesis is to apply s-SNIM to model systems for optimizing the sensitivity, checking the biological compatibility and the limitations of this new technique.
SAMs have proven to be very well-suited for this task due to the large number of different organothiols available (easily interchangeable absorbing tail groups), due to their
ease of patterning (µCP, dip-pen, nanografting, etc.) and due to their compatibilities
with gold substrates which are offering an extra enhancement of the near-field signal.
The self-assembly process of thiolates on gold is well documented into the literature
therefore allowing the growth of well defined SAMs on gold which have an enormous
potential as model system for optimizing and understanding the contrast mechanism
driving near-field microscopy. In the long-term the understanding of the contrast generated by patterned model surfaces will help studying unknown or more complex samples
(e.g. living system, cell membranes).
Extremely powerful is the methodology of fabricating in close proximity laterally structured distinct chemical functionalities. Per se these different chemical functionalities
defining and controlling interfacial reactivity patterns are already interesting especially
when the patterns are in the nanoscale (e.g. DNA hybridization nanosensors). Another
very important aspect is that the patterning always can be used for defining reference
areas avoiding the need for gold beads which are always bearing the risk of contaminating the near-field contrast with topography artefacts.
In this thesis the potential of s-SNIM as a very sensitive, label free and non-invasive
technique for obtaining IR spectroscopical information with nanometer resolution is established.
The ability of s-SNIM to provide infrared spectroscopical information on thin organic
films is first demonstrated with microstructured monolayers of 1-octadecanethiolate
(ODT) and a biotinylated alkylthiolate (BAT). At a wavelength of 5.85 µm a lateral
resolution of ∼90 nm was achieved comparable with a diffraction limited resolution of
λ/60. The detection limit achieved was 5 x 10−20 mol corresponding to 27 attogram or
30.000 molecules of BAT.
In the DNA chapter the s-SNIM was applied to a biological system showing in a proof
of concept the high potential of s-SNIM for a new generation of label free nanoscale
biosensor platforms. Nanografting of DNA and IR-nanoscopy were combined to study
DNA hybridization in nanostructured DNA SAMs.
Further enhancement of s-SNIM sensitivity can be achieved with IR labels such as metalorganic compounds. These dyes are especially with regard to application of s-SNIM
in life sciences very interesting (e.g. cell studies, biosensors). IR dyes enable enhanced
sensitivity because their resonance fall into spectral regions around 2000 cm−1 where no
natural absorption is seen.
2
2. Theory of self-assembled
monolayers (SAMs) on gold
Contents
2.1. Molecular self-assembly . . . . . . . . . . . . . . . . . . . . . .
4
2.2. Brief historical introduction to SAMs . . . . . . . . . . . . .
4
2.3. Basic theory of thiolate-gold SAMs
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2.3.1. Gold substrates . . . . . . . . . . . . . . . . . . . . . . . . . .
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2.3.1.1. Physical vapor deposition (PVD) of gold . . . . . .
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2.3.1.2. Template stripped gold . . . . . . . . . . . . . . . .
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2.3.1.3. Cleaning of gold substrates . . . . . . . . . . . . . .
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2.3.2. Characteristics of alkanethiolates on gold . . . . . . . . . . .
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2.3.2.1. Structure of alkanethiolates on gold . . . . . . . . .
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2.3.2.2. Formation of alkanethiolates on gold . . . . . . . . .
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2.3.2.3. Stability of alkanethiolates on gold . . . . . . . . . .
11
2.3.3. Analytical techniques for characterization of SAMs . . . . . .
12
2.4. DNA SAMs - a special case . . . . . . . . . . . . . . . . . . .
13
2.4.1. Basics on deoxyribonucleic acid (DNA) . . . . . . . . . . . .
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2.4.1.1. Molecular structure . . . . . . . . . . . . . . . . . .
13
2.4.1.2. Hybridization . . . . . . . . . . . . . . . . . . . . . .
15
2.4.1.3. Secondary structure . . . . . . . . . . . . . . . . . .
16
2.4.2. Structure of DNA SAMs . . . . . . . . . . . . . . . . . . . . .
17
2.4.3. Hybridization of immobilized DNA . . . . . . . . . . . . . . .
19
2.4.4. Techniques for label-free detection of hybridization . . . . . .
19
3
Chapter 2. Theory of self-assembled monolayers (SAMs) on gold
2.1. Molecular self-assembly
Molecular self-assembly is a natural spontaneous organization process of molecules. Two
types of molecular self-assembly are known:
i) intramolecular self-assembly (commonly called folding) occurs for instance when a
protein chain folds from its random coil conformation into a well-defined tertiary
stable structure.
ii) intermolecular self-assembly means the spontaneous organization of two or more
molecules. An example is the assembly of two complementary single stranded
strands of DNA into a double helix or the formation of micelles, liposomes or
bilayers from single lipids caused by hydrophilic and hydrophobic interactions.
The driving force for molecular self-assembly is a decrease in free energy ∆G and eventually reaching a state of minimized energy. This process is directed through noncovalent
interactions (e.g. hydrogen bonding, hydrophobic forces, van der Waals interactions,
pi-pi interactions, electrostatic effects or metal coordination).
Self-assembly is referred to as bottom-up technique in contrast to top-down techniques
such as lithography.
2.2. Brief historical introduction to SAMs
Intermolecular self-assembly at interfaces can result in molecular films with a thickness of
one molecule. Those films are called self-assembled monolayers. The formation of SAMs
can occur at different interfaces. Generally molecules arranging into a self-assembled
monolayer at solid surfaces can be separated into three parts: a head or anchor group,
which interacts with a substrate, a spacer or linker, and a tail or terminal functional
group which determines the surface properties (Fig. 2.1 and Table 2.1).
The first self-assembled films with a thickness of only one molecule were detected in 1917
by Langmuir [2] when he studied the spreading of amphiphiles on water (Fig. 2.2 A).
Figure 2.1.: Schematic diagram of an ideal, single-crystalline SAM on a support
4
2.2. Brief historical introduction to SAMs
part
head or
anchor group
spacer
or
linker
-
terminal functional or
tail group
properties
provides chemical functionality with
specific affinity for a substrate
organic interphase(1-3 nm)
provides well-defined thickness
acts as a physical barrier
alters electronic conductivity
and local optical properties
determines surface properties
presents chemical functional group
examples
−SH on Au, Ag, Cu;
−SiCl3 on SiO2
−(CH2 )n −
−C6 H4 −
−CH3 , −OH, −COOH,
−(O−CH2 −CH2 )x −OH
mit x=3-6, −NH2
Table 2.1.: Properties of the three principal components of SAM forming molecules.
Different combinations of head groups and substrates are listed in [1].
In 1935 Katharine Blodgett was the first who transferred monomolecular films from an
air-water interface to a solid support [3]. The Langmuir-Blodgett technique is still today an often used method. A solid support is dipped into an aqueous solution with a
self-assembled monolayer at the air-water interface. When pulling the support carefully
out of the solution the molecular monolayer is transferred onto the surface of the solid
support (Fig. 2.2 B). Repetitive dipping of the solid support into the water trough results in the formation of multilayers. A drawback of the Langmuir-Blodgett method is
the thermodynamic instability of the films. Temperature variations or exposure to many
solvents easily ruin the 2D structure of the Langmuir-Blodgett films.
A second type of monolayer deposition was developed in 1946 by W.A. Zisman and
coworkers [4]. In this method the monolayers spontaneously assembled on the surface of
a metal substrate. Therefore the metal was immersed into a solution containing a surfactant (Zisman used platinum substrates and alkylamines as surfactants) and the system
was given some time to equilibrate (Fig. 2.2 D). Since then different adsorbate/substrate
combinations forming SAMs have been found. Among them alkyltrichlorosilanes on glass
[5] and fatty acids on metal oxide surfaces [6]. In 1983 Ralph Nuzzo and David Allara
discovered the formation of SAMs from diluted solutions of alkylthiols and disulfides on
gold [7]. Organosulfur derivatives (thiols, disulfides, sulfides) on gold are even today the
most studied and most popular systems due to their ease of use.
Due to the great variety of different substrate-head group combinations and the wide
choice of terminal functional groups SAMs provide a powerful tool to control, change and
tailor interfacial properties of surfaces. Their great flexibility has led to a broad range of
applications in different areas. Even now-a-days the field of applications is still growing
5
Figure 2.2.: Schemes of different techniques for monolayer deposition on solid supports.
A) Langmuir monolayer B) Langmuir-Blodgett films C) Monolayer deposition from solution D) Monolayer deposition from gas phase
and getting more diverse. In life sciences SAMs can serve as excellent simplified model
systems for complex biological interfaces like for instance the cell membrane. Such
artificial surfaces with biological functionality allow detailed studies on fundamental
aspects of biological systems like protein adsorption or cell adhesion. Furthermore SAMs
can be used to tailor biological relevant surfaces that require stringent control of surface
properties like implants or biosensors.
2.3. Basic theory of thiolate-gold SAMs
Since the SAMs used in the experimental part of this thesis are based on a thiolate-gold
bonding only the basic theory about this SAM system is outlined. For more detailed
information the reader is referred to recently published reviews on the self-assembly of
thiolates [1, 8]
2.3.1. Gold substrates
Gold has some characteristics that make it well-suited as a substrate for studying SAMs.
• gold surfaces are easy to prepare and are commercial available
• gold is inert towards oxidization at temperatures below its melting temperature, it
does not react with atmospheric oxygen, and it does not react with most chemicals.
For this reasons gold is easy to handle and does not require high vacuum conditions.
6
2.3.1. Gold substrates
• gold has a strong specific interaction with sulfur [9] that allows to form monolayers
in the presence of many other functional groups
• gold surfaces are common substrates for a number of analytical techniques e.g.
infrared reflection absorption spectroscopy (IRRAS), electrochemical techniques,
surface plasmon resonance (SPR) or scanning near-field infrared microscopy (SNIM)
• gold shows no toxicity toward cells and is therefore well-suited for studies and
applications of SAMs as interface in biological research.
2.3.1.1. Physical vapor deposition (PVD) of gold
Several methods for the deposition of gold films on silica surfaces have been developed.
The most commonly used process is physical vapor deposition which is a general term
for a variety of techniques where thin films grow by condensation of vaporized material.
In PVD most techniques differ in the way of material/metal vaporization. The gold
substrates used within this thesis are fabricated by evaporation or sputter deposition.
For evaporation deposition the gold (or another material/metal) is electrically heated
to a high vapor pressure until it sublimes or evaporates. Upon sputter deposition high
velocity Ar− ions from an Argon plasma bombard a gold cathode. Due to momentum
transfer cathode surface material is sputtered and the ejected gold atoms condense on a
substrate forming a thin gold film.
Gold such as most metals (especially noble metals) has poor adhesion on silicon or silicon
oxide due to the different lattice parameter of both materials. To promote the adhesion
a primer layer (typically 1-5 nm) of titanium or chromium is deposited on the silica
surface followed by the deposition of gold (typically 50-200 nm). This adhesion layer
helps to bond the gold to the substrate and prevents peeling off the gold layer during
subsequent processing and chemical treatments. Long storage of such substrates is not
recommended because the primers tend to diffuse through the overlying gold film to the
surface over time [10]. If an adhesion layer is not desirable (e.g. for template stripping),
gold can be deposited directly onto the substrate.
Gold films on silica surfaces are polycrystalline, which means that they consist of many
small, single-crystalline grains. The grain size can vary depending on the substrate and
the deposition parameters (e.g. temperature of the substrate during deposition, angle
of incidence of vapor on the substrate, deposition rate, etc.). Generally grains of gold
on silica range in size from ∼10 to 1000 nm. Since many properties, e.g. mechanical,
electronic, magnetic, optical or chemical are influenced and often even dominated by the
specific grain structure it is desirable to manipulate grain sizes after deposition, e.g by
heat or stress treatments.
For SPM gold surfaces with very large and flat areas are required. Since typical organic
molecules for SAMs have lengths between 1-5 nm similar to the roughness of the gold the
roughness is an important factor when characterizing SAM properties and distinguishing
molecules with small height differences within mixed or lateral structured SAMs. Surface
roughness is also an important issue in SPM based nanolithography [11].
Hence, several procedures for the preparation of flat gold surfaces have been reported
7
including flame annealing, chemical etching [12], and direct evaporation of gold onto flat
surfaces such as mica or molecular adhesion monolayers [13]. A practical drawback of
these techniques is the requirement of gold deposition immediately before use, which is
very time-consuming for routine use.
2.3.1.2. Template stripped gold
A widely used method for preparation of flat gold substrates that allows for the rapid exposure of large areas of ”fresh” gold surfaces on demand is template stripping. Template
stripping was first introduced by G. Semenza and co-workers in 1993 [14] and is able to
produce large areas (up to several µm) of smooth ultraflat gold (∼2-3 Å). Generally a
silica-based substrate is glued on top of a gold layer deposited directly on a mica plate
(template) without any adhesion layer. After curing, the gold is either mechanically
separated from the template (mechanical stripping) or with the aid of a penetrating
solvent (chemical stripping). The exposed template-stripped gold (TSG) surface (i.e.
the very first atom layer of gold having deposited onto the template) is nearly as flat as
the template itself and is a replica of the mica surface.
Beneficial is that the precursors (template/gold/support ”sandwich”) can be prepared in
bulk and stored up to several months without loss of quality [14]. This means ”fresh” gold
surfaces can be quickly obtained immediately before use only by opening a precursor.
However, template-stripped ultraflat gold surfaces are not easy to make. Stripping the
mica off the gold surface is not always complete and mica residues still remain attached
to the gold. Therefore the template stripping method has been expanded to utilize
templating surfaces other than mica [15, 16]. However, the mechanical stress during
the stripping procedure can result in gold surfaces with defects. Another shortcoming
is that depending on the type of glue used for preparation, exposure to some types of
solvents can cause swelling creating roughness and disruptions on the gold surface [17]
or the glue can dissolve. To overcome those problems studies on solvent resistance of
different glues [17] and non-glue based strategies were reported [18].
Criteria for selecting the type of substrate and method of preparation are dependent
on the application for which the substrate is used and on the laboratory equipment
available.
2.3.1.3. Cleaning of gold substrates
An essential requirement for high quality alkylthiolate monolayers is a clean Au substrate. Gold is chemically nonreactive, but is instantaneously covered with a reversible
physisorbed layer of water, hydrocarbons, and other organic compounds under ambient
laboratory conditions ([19] and references therein). Straight chain alkanethiols are able
to replace weakly adsorbed contaminations during SAM formation especially upon prolonged incubation times but other thiols with large bulky head groups or bad ordering
properties may not be able to fully remove the contaminants resulting in non-ideal layer
formation. Therefore, it is recommended to clean gold substrates immediately prior to
8
2.3.2. Characteristics of alkanethiolates on gold
use when the gold was not deposited freshly (within 1 hour) or stored in a special way
(e.g. template stripping precursors).
There are several methods available for cleaning gold substrates such as oxygen plasma
treatment or strongly oxidizing chemicals. The most common procedure uses piranha
solution, which is a strong oxidant and removes most organic contaminants. Piranha
solution typically consists of a 3:1 (v/v) mixture of 96% H2 SO4 and 30% H2 O2 (other
protocols use 7:3 or 4:1 mixtures) and should be used freshly prepared. Gold substrates
should not be left too long in the solution, since the roughness of the gold surface increases upon prolonged exposure. Cleaning usually requires 5-15 min. A clean gold
surface is hydrophilic [20, 21] and completely wetted by water. The gold layer may even
peel off, especially if no adhesion layer between the gold and the substrate is present.
2.3.2.1. Structure of alkanethiolates on gold
Upon exposure of thiols towards a gold surface the hydrogen from the thiol group (-SH)
dissociates and a thiolate-gold bond is formed:
1
X−(CH2 )n −S−H + Au(0)m → X−(CH2 )n −S− Au(I) · Au(0)m−1 + H2 ↑
2
(2.1)
X = terminal group
The thiolate-gold bond is relatively strong (∆H0 ∼160 kJ/mol or ∼40 kcal/mol [22, 1])
and thus the resulting monolayers are quite stable. This stability is highly preferable for
applications in biological surface science since it allows to study interface interactions
under physiological conditions. Details on the nature of the metal-sulfur bond and it
spatial arrangement of the sulfur groups on the underlying gold lattice are still controversial.
√ √ However, the high coverage thiol phase on Au (111) is generally described as a
( 3x 3)R30◦ (R=rotated) overlayer [23, 24, 25, 26]. The spacing between adjacent sulfur atoms in this structure is approximately 4.99 Å with a footprint of 21.4 Å2 /molecule
Figure 2.3.: Schematic diagram of an ideal, single-crystalline SAM of alkylthiolates
supported on a gold surface
9
[27]. These figures represent the upper density limit of the SAM. For the alkyl chains of
a thiolate SAM on gold a separation distance smaller than the sulfur-sulfur distance is
found (4.24 Å vs. 4.99 Å) resulting in a tilting of the chains of about 27◦ with respect
to the surface normal (Fig. 2.4). For this orientation the total free energy of the system
is minimized by optimizing the interchain van der Waals interaction (each methylene
group contributes ∼1.5 kcal/mol of stabilization to the SAM [28]) and the sulfur-gold
interaction. Furthermore the alkyl chains are fully extended in a nearly all-trans configuration of the carbon-carbon bonds. The twist angle, β, which describes the rotation of
the CCC bond plane relative to the plane of the surface normal and the tilted chain is
about ∼52◦ (Fig. 2.4).
Figure 2.4.: Schematic view of the orientation of a single, long-chain thiol molecule
adsorbed on gold. The tilt angle α is with respect to the surface normal, whereas the
twist angle β is with respect to a plane established by the chain axis and the surface
normal vectors.
The packing density of the chains is determined by the spacing of the sulfur atoms and
is similar to that of polymethylene chains in bulk, crystalline n-alkanes. However, for
the case of short chain length (n < 8) a loss of film organization (loose packing and
increasing disorder) was reported [29, 19].
Order and orientation of the terminal group of an ω-substituted alkanethiolate dominates the properties and interaction of the interface between the SAM and a contacting
phase. Unfortunately, the tail group does not only introduce the desired surface termination but often is also the reason for an imperfect self-assembly, e.g. disordered
SAMs. In general, the tendency for a monolayer to deviate from well-ordered structures and to exhibit structural disorder and defects depends on the matching between
the van der Walls radii of the tail group and the head group and their fitting with the
substrate lattice parameters. In order to overcome this aspect highly charged or bulky
tail group containing thiols are often accommodated by ”diluting” them in a monolayer
of shorter chain thiols. Dilution also ensures the functionality of bulky head groups e.g.
10
biotinylated alkylthiols which recognize streptavidin [30].
2.3.2.2. Formation of alkanethiolates on gold
SAM formation can be performed either from gas or vapor phase or from solution
(Fig. 2.2 C, D). In most cases adsorption from solution is favored since it requires no
ultra high vacuum (UHV) conditions. Scanning probe microscopy (SPM) and diffraction studies reveal that SAM formation from solution occurs in two steps: initially at
low coverage the molecules attach to the gold with their hydrocarbon chains oriented
parallel to the surface and lying down on the surface (striped phase). With increasing
coverage thiols from solution induce a lateral pressure causing the molecules to raise
up into a standing-up configuration and form a crystalline or semicrystalline structure
(crystalline phase).
The rate of SAM formation is influenced by many factors such as temperature, solvent,
concentration, chain length of the adsorbate, and cleanliness of the substrate. Experimental conditions must be established for each new system studied. The formation
kinetics of a monolayer is biphasic. The initial adsorption is rapid and diffusion controlled resulting in an imperfect monolayer (a clean gold surface placed in a 1 mM
solution of ODT in ethanol is hydrophobic after about 2 s). It is followed by a slower
period lasting several hours where the molecules orient, displace contaminants, reduce
defects by lateral diffusion and enhance packing ((re)-crystallization process).
Figure 2.5.: SAM growth in solution
2.3.2.3. Stability of alkanethiolates on gold
Monolayers of alkanethiolates on gold appear to be stable in air or in contact with liquid
water, salt solutions or ethanol at room temperature. They have been used for studies
of protein adsorption and cell adhesion in aqueous media over periods of several days.
Upon heating to temperatures over 70◦ C the molecules desorb [19]. Damages of the
monolayer can be caused by chemicals that include halogens (I2 , Br2 ), strong oxidizing
agents (peroxides, ozone) or chemicals that attack the gold film (e.g. aqua regia) or the
adhesion layer (concentrated HCl).
However, SAMs of alkanethiolates are stable barriers protecting the underlying gold from
dissolution in highly corrosive etchants, such as aqueous CN− /O2 [31].
11
2.3.3. Analytical techniques for characterization of SAMs
Since monolayers are used in a broad range of applications a basic knowledge on their
formation, final structure and interaction with other molecules is essential. From an
analytical point of view especially SAMs of thiolates on gold are very suitable for such
studies since gold surfaces are compatible with many techniques: (i) thin films of gold
(<100 Å) are optical transparent, gold is electrical conductive, and SAMs of thiolates
on gold are very stable. An overview of different analytical techniques used for characterization of SAMs is given in Table 2.2.
Analytical technique
General
Contact Angle
QCM/SAW
Optical
IR spectroscopy
UV-Vis absorbance
Fluorescence spectroscopy
Ellipsometry
SPR
Vacuum
XPS
AES
SIMS
Microscopy
AFM
STM
Electrochemical
Cyclic voltammetry
Impedance spectroscopy
structural information
hydrophobicity, order
changes in mass, kinetics
functional groups, molecular orientation
density of adsorbates
density of adsorbates
layer thickness, refractive index
layer thickness, kinetics
elemental composition
elemental composition
molecular mass of adsorbate (+ fragments)
molecular packing
molecular packing
thickness, order/defects
thickness, order/defects
Table 2.2.: Analytical techniques for monolayer characterization according to Ref [32].
QCM = quartz crystal microbalance, SAW = surface acoustic wave, SPR = surface plasmon resonance, XPS = X-ray photoelectron spectroscopy, AES = Auger electron spectroscopy, SIMS = secondary ion mass spectrometry, AFM = atomic force microscopy,
STM = scanning tunneling microscopy.
12
2.4. DNA SAMs - a special case
2.4. DNA SAMs - a special case
2.4.1. Basics on deoxyribonucleic acid (DNA)
2.4.1.1. Molecular structure
DNA is a polymer consisting of structural units called nucleotides. Each nucleotide
has three compounds (Fig. 2.6): a base, a pentose molecule (in the case of DNA 2deoxyribose), and a phosphate group. In general DNA contains four different bases:
adenine (A), cytosine (C), guanine (G), and thymine (T) which can be classified into
two types: adenine and guanine are heterocyclic molecules called purines, while thymine
and cytosine are homocyclic molecules called pyrimidines.
Figure 2.6.: Schematic structure of a nucleotide consisting of a phosphate group (blue
circle), a sugar molecule in the case of DNA 2-deoxyribose (orange circle), and a base.
The four bases found in DNA are displayed on the right: thymine (T, turquoise), adenine
(A, red), guanine (G, green), and cytosine (C, violet). The homocyclic bases T and C
are called pyrimidines and the heterocyclic bases A and G are called purines.
In a polynucleotide the nucleotides are linked by the phosphate group which connects
two nucleotide sugars by forming a phosphodiester bond (Fig. 2.7). This results in a
DNA strand backbone with alternating phosphate groups and sugar molecules which is
due to the free charges in the phosphate groups negatively charged.
Each polynucleotide has two different ends (Fig. 2.7): on the one end it has a terminal
phosphate group referred to as 5’ (five prime)-end and on the other end it has a terminal
hydroxyl group referred to as 3’ (three prime)-end. By definition a DNA strand starts
at the 5’-end. This is especially important when writing the order of the bases along the
DNA strand that is the sequence of the DNA. The term used for the length of a DNA
13
Figure 2.7.: Schematic structure of a 4 bp DNA double strand. Nucleotides are linked
by phosphodiester bonds (gray box). Thymine (T, turquoise) and adenine (A, red) form
two hydrogen bonds, guanine (G, green) and cytosine (C, violet) form three hydrogen
bonds. The dashed lines between the bases display the hydrogen bonds.
sequence is x-mer or x bp (base pairs), e.g. for a sequence consisting of 25 bases: 25-mer
or 25 bp. Short segments of DNA typically up to 30 bases are called oligonucleotides.
In nature the size and composition of the DNA determine the form and function of a
resulting organism. Relative simple organisms such as bacteria have genomes1 of approximately 1-5 million bases while the human genome has approximately 3000 million bases.
In living organisms DNA exists as double strand in which two nucleotides held together
by hydrogen bonds between their bases and form a helical structure (Fig. 2.8) Only
complementary bases form hydrogen bonds such that A interacts with T forming two
hydrogen bonds and G interacts with C forming three hydrogen bonds (Watson-Crick
rule)(Fig. 2.7). Due to this complementary base pairing DNA binding is very specific.
In a double-strand the two polynucleotides are oriented antiparallel which means that
their directions are opposite (Fig. 2.7). Additionally to the hydrogen bonds between the
1
genome: complete genetic sequence on one set of chromosomes
14
2.4.1. Basics on deoxyribonucleic acid (DNA)
complementary bases the double helix is stabilized by hydrophobic interactions between
the bases and pi stacking. Within the helix the aromatic rings of the bases are positioned
nearly perpendicular to the backbone of the strands. Thus, the faces of the aromatic
rings are arranged parallel to each other allowing the bases to interact through the pi
orbitals.
The helical structure exists in different conformations: A-, B-, and Z-DNA. A- and BDNA are right-handed spirals whereas the spiral of the A form is wider than that of the
B-form. In living organisms the B-form is most common and the A-form occurs under
non-physiological conditions in dehydrated samples. The Z-DNA is a left-handed spiral
that can be found for instance in DNA segments where the bases have been chemically
modified by methylation. More structural properties of this DNA forms are given in
Table 2.3.
Figure 2.8.: Schematic drawing of a DNA helix corresponding to the DNA sequence used
in this thesis.
structural properties
A-DNA
helix sense
right-handed
diameter [nm]
∼2.6
base pairs per turn
10.7
rise per turn [nm]
3.4
rise per base [nm]
0.29
major groove
tight, low
minor groove
broad, flat
B-DNA
right-handed
∼2.0
10.0
3.4
0.34
broad, low
tight, low
Z-DNA
left-handed
∼1.8
12
3.4
4.4
tight, low
flat
Table 2.3.: Comparison geometries of the most common DNA forms
2.4.1.2. Hybridization
The process by which two complementary polynucleotide chains form a stable double
helix is known as hybridization. Hybridization reactions can occur between two complementary DNA molecules in solution or between a molecule in solution and a complementary molecule immobilized on a solid support. The driving forces of hybridization
are differences in the thermodynamical equilibrium between double- and single strands.
Many factors influence the hybridization process. On the one hand the properties of
15
the DNA sequence itself play an important role, such as sequence composition, strand
length, and secondary structure. On the other hand external conditions such as temperature and salt concentration affect the double-strand formation.
The reversal process of hybridization, the separation of a double helix into two single
DNA strands, is known as melting or denaturation of DNA. Denaturation of nucleic
acids occurs with extreme pH, low ionic strength, heat, and chemicals such as urea.
Usually denaturation is achieved by heat treatment. The temperature at which 50%
of the double stranded DNA have been dissociated into single strands is defined as the
melting temperature Tm of DNA. The melting temperature of dsDNA, and hence its
stability, depends on several factors, including the DNA sequence, solvent, concentrations of ions in solution and the pH. Tm is characteristic for a given DNA sequence and
depends on the percentage of GC basepairs. With rising GC content Tm increases due to
the higher energy needed for the breaking of three hydrogen bonds between guanine and
cytosine compared to the energy needed for breaking of two hydrogen bonds between
adenine and thymine. High ion concentrations shift the melting temperatures to higher
values. The most convenient way of monitoring thermal denaturation of DNA is by
its ultraviolet (UV) absorbance which is caused from the aromatic nature of the bases.
When DNA denatures, its UV absorbance increases by about ∼40 % at all wavelengths
whereby the shape of the absorbance curve does not change [33]. This phenomenon,
known as the hyperchromic effect, results from the disruption of the electronic interactions among nearby bases. The heterocyclic rings of the bases absorb more if they are
not connected by hydrogen bonds and are not piled up. DNA’s hyperchromic shift upon
thermal denaturation is usually monitored at 260 nm. Because hydrogen bonds are no
covalent bonds the melting and hybridization of DNA is a reversible process.
Hybridization is the biochemical process on which the entire DNA microarray industry
is based. In biochemical assays the DNA in question is called target and the complementary DNA (probe). In DNA arrays generally the probe is immobilized on a surface.
For surface-immobilized probes the thermodynamic equilibrium conditions changed compared to probes in solution. The hybridization can be kinetically or sterically embarrassed or inaccessible for some sequences leading to low hybridization efficiencies and/or
excessively long incubation times.
2.4.1.3. Secondary structure
The primary structure of a polynucleotide is given by its base sequence. Single-stranded
DNA that contains self-complementary bases can form intramolecular base pairs resulting in the formation of a secondary structure. A well-known secondary structure of
DNA is a hairpin (Fig. 2.9). It forms when two complementary domains are separated
by some non complementary bases. After base pairing of the complementary domains
the non complementary bases form a loop.
Secondary structure is a critical issue affecting the hybridization of polynucleotides.
Studies on energetic comparable random coil (polynucleotides without intramolecular
16
2.4.2. Structure of DNA SAMs
Figure 2.9.: Schematic drawing of a DNA with a hairpin secondary structure.
base pairs) and secondary structured DNA sequences demonstrate that at high temperatures the activation energies of DNA hybridization are negative and independent of
secondary structure. But with decreasing temperature the intramolecular base pairs in
secondary structured DNA stabilize the single-strand conformation resulting in a much
higher hybridization activation energy than for the random-coil DNA [34, 35]. Hindered
hybridization due to secondary structures is a critical issue in DNA chip technology. Secondary structures in either probe or target strand can impair or prevent hybridization
resulting in false results (negative hybridization) during read-out of the signal intensities from the different microarray spots. In order to avoid such effects hybridization
is commonly carried out at increased temperatures and much effort is given on probe
or target design. Different algorithms have been designed to predict secondary structures of polynucleotides under various solution conditions. A very popular software for
thermodynamic predictions on secondary structure and DNA hybridization is Mfold by
Zuker [36].
2.4.2. Structure of DNA SAMs
DNA can be modified with a thiol group that is linked to one end of the strand via a
carbon chain. Commonly the thiol group is attached to the 5’-end using a chain of six
carbon atoms as spacer (Fig. 2.10).
Figure 2.10.: Thiol group with a six carbon linker attached to the 5´-end of a DNA
molecule
SAMs formed from thiol modified DNA molecules do not order as well as alkylthiolate
monolayers. Four aspects play a crucial role for DNA adsorption upon immobilization:
17
(i) unspecific interactions between gold and DNA:
Upon adsorption not only the thiol group interacts with the gold but also the amine
moieties in the DNA bases can chemisorb weakly on gold [37, 38] and electrostatic
interactions between gold and the nucleotides occur. A common method to displace non-thiol interactions of the DNA with the gold surface is backfilling with
6-mercapto-1-hexanol (MCH)2 [39]. Unfortunately post-treatment with MCH does
not completely replace non-thiol DNA interactions [40] but it was recently reported
that co-immobilization of thiolated DNA and MCH results in better ordered SAMs
than achieved with the posttreatment method [41].
(ii) electrostatic repulsion between negatively charged DNA strands:
The negative charges in the DNA backbone cause electrostatic repulsion among
different strands. In high ionic strength solutions the negatively charged backbones are effectively shielded. Consequently faster adsorption kinetics and higher
molecular densities can be reached in high salt solutions than in low salt solutions [39, 42]. Furthermore even the cationic nature influences the immobilization
efficiency, which is dramatically increased in divalent salt solutions [43].
(iii) flexibility of the DNA strands:
Persistence length [44] is a parameter which quantifies the bending flexibility of
polymers:
l ∼ B/kB T
(2.2)
l = persistence length, B = bending elastic constant, kB = Boltzmann’s constant, T =
temperature
It describes the distance over which a polymer behaves like a rigid rod. Singlestranded DNA has a persistence length of only about 1 nm (a single base has a
length of 0.4 nm [45]). Thus, most immobilized polynucleotides are much longer
than their persistence length and consequently form tethered random coils instead
of orienting perpendicular to the surface.
In case of double-stranded DNA the persistence length is dramatically increased
to about 45 nm [46] (∼130 bp, base length double-stranded B-form DNA: 0.34
nm). Consequently DNA helices shorter than 130 bp behave like a rigid rod.
Comparison studies between immobilization of ss-DNA and ds-DNA result in a
lower final coverage and faster adsorption kinetics for the ds-DNA. However, no
differences in hybridization kinetics were found when comparing the same probe
densities [42].
(iv) DNA length:
Thiolated DNA sequences shorter than 24 bases tend to adsorb mainly via the thiol
2
MCH was selected for three reasons: (i) DNA does interact with MCH, (ii) MCH is soluble in aqueous
solution and therefore compatible with biological applications, and (iii) MCH has the same length
as the 6-carbon chain linker in the thiolated DNA avoiding interferences with the complementary
strand during hybridization.
18
2.4.3. Hybridization of immobilized DNA
group to the gold whereas longer strands attach with multiple sites to the gold
resulting in a less ordered arrangement and a length dependent surface coverage
[40].
However, DNA surface density can be roughly controlled over the amount of time that
gold is exposed to thiolated DNA solution but to this end all above mentioned aspects
have to be considered. Under conditions of 1 M salt (NaCl or KH2 PO4 ) the coverage
increases strongly to approximately 50% of the maximal coverage within 15 minutes.
Then the adsorption kinetic slows down and after 2 hours approximately 80% of the
maximal coverage (after 20 h) is achieved. Further exposure results in little additional
adsorption due to saturation of the surface [39, 42].
2.4.3. Hybridization of immobilized DNA
It is known that a solid support affects DNA hybridization on surfaces but it is difficult
to compare measurements of hybridization in solution and on surfaces, among others
due to the large differences in DNA concentration and effective ionic strength in these
two environments. Significant differences between solution and surface hybridization
kinetics and thermodynamics were recently reported by Georgiadis and coworkers [35].
In comparison to hybridization efficiency in solution surface hybridization reaches only
15-25% efficiency under conditions of target saturation and high probe densities of 4.5 to
6.8 molecules/cm2 . Moreover surface hybridization kinetics are decreased by a factor of
20- to 40-fold compared to solution phase kinetics. These results are mainly attributed
to steric and electrostatic hinderance caused by the confinement of the immobilized DNA
as well as conformational restrictions of the immobilized strands.
Previous studies on immobilized DNA show also an inhibited hybridization efficiency at
high probe densities [39, 42]. However, at low probe densities (≤3x1012 molecules/cm2
[42]) hybridization efficiencies of up to ∼100% have been reported [39]. Heating of the
probe film prior or upon hybridization significantly increases hybridization efficiency
in the low density regime of about 20% compared to a not heated surface. However,
hybridization efficiency at very high probe densities is not affected by heat treatment.
At low probe densities the probe-target capture follows a Langmuir-like kinetic. It is
drastically slowed down at high probe densities [42].
2.4.4. Techniques for label-free detection of hybridization
Most hybridization detection approaches rely on the labeling of samples with fluorophores, radioactive, or electrochemical tags. This allows for very sensitive detection
but can be time-consuming and expensive. Beside these commonly used methods labelfree alternatives for detection of hybridization are listed here:
• quartz crystal microbalance (QCM):
detects changes in mass during hybridization by measuring the change in frequency
of a quartz crystal resonator on which the ss-DNA is immobilized
19
• surface plasmon resonance (SPR):
measures changes in the refractive index at a metal-adsorbate interface which
changes upon hybridization
• atomic force microscopy (AFM):
monitors topographical and elasticity changes of DNA nanofeatures upon hybridization (for details see chapter 11.4.1 (page 139)
• infrared spectroscopy:
records changes in vibrational modes of DNA bases during hybridization (for details see chapter 11.3 (page 128)
• electrochemical impedance spectroscopy [41]:
detects changes in the charge transfer resistance upon hybridization due to electrostatic repulsion from the negatively charged DNA (a certain threshold probe
density is required)
• time-resolved THz spectroscopy [47]:
measures differences in the DNA binding state of DNA due to changes in its
dielectric properties (e.g. interbackbone excitations of DNA molecules)
Most of these techniques are capable of detecting DNA quantitatively but their relatively
large detection areas are limiting their detection sensitivity.
20
3. Theory of lithographic techniques to
laterally structure SAMs
Contents
3.1. Microcontact printing (µCP) . . . . . . . . . . . . . . . . . .
22
3.2. Scanning probe lithography (SPL) . . . . . . . . . . . . . . .
26
3.2.1. Nanografting . . . . . . . . . . . . . . . . . . . . . . . . . . .
27
3.2.1.1. Nanografting of biological molecules . . . . . . . . .
29
21
Chapter 3. Theory of lithographic techniques to laterally structure SAMs
Molecular patterning of surfaces plays an important role for fundamental cell biology,
tissue engineering and biosensor technology. For instance it is relevant for studying
the interaction of living cells with surfaces and to control their growth and spatial
arrangement. In pharmacological and biochemical research different kinds of assays
(e.g. microarrays) require control over spatial distribution of biomolecules (e.g. proteins
or nucleic acids) adsorbed on surfaces.
In this chapter two different techniques for lateral structuring of SAMs are introduced.
In the first part microcontact printing which belongs to the class of soft lithography
methods is outlined. The second part introduces scanning probe lithography with a
main focus on nanografting. Nanografting can help to address fundamental questions
about protein binding or DNA hybridization at the molecular scale.
3.1. Microcontact printing (µCP)
Microcontact printing is a soft lithography technique for lateral structuring of SAMs
that was developed in 1993 at the Harvard University by Whitesides and coworkers [48].
The technique works analogous to printing ink with a rubber stamp on paper. But in
case of µCP the inks are solutions of thiolated molecules and the paper is substituted
by a gold surface. If the size of the printed features is in the nanometer scale the technique is also called nanocontact printing. Two excellent reviews on soft lithography are
published by the group of G. M. Whitesides [49, 50].
For fabricating a stamp a rigid substrate (master) having a desired relief is necessary.
Such masters can be for instance commercially available diffraction gratings, TEM grids,
sieves or AFM calibration standards. Master with other desired patterns can be fabricated by conventional lithographic techniques like photolithography, e-beam writing or
anisotropic chemical etching. Stamp fabrication starts with casting a freshly mixed and
viscous prepolymer onto the master. After curing, the elastomer is gently peeled off and
an elastomeric stamp with a negative template relief is fabricated.
The most common stamp material is poly(dimethylsiloxane) (PDMS) which is a soft,
chemically cross-linked silicon rubber that is commercially available as Sylgard 184 (Dow
Corning Corporation). It is well-suited for µCP for following reasons: it is
• nontoxic
• inert towards most chemicals
• compatible with a wide range of organic and organometallic molecules
• not hydroscopic
• elastomeric (Young´s modulus approximately 1.8 MPa) allowing the stamp to
adapt even to nonplanar surfaces
22
Figure 3.1.: Polymerization reaction of PDMS
• has a low surface energy (γ = 21.6 dyn/cm2 ) that allows an easy removing of the
stamp from most surfaces and a relative resistance of the stamp surface against
contaminations by adsorption or reaction with ”ink” molecules and dust particles.
The polymerization reaction of PDMS is shown in Figure 3.1.
Stamp loading (”inking”) can be performed in different ways mainly depending on the
available amount of ink solution. The most common procedure is to immerse the stamp
in a solution of the ink molecule (e.g. an ethanolic solution of alkylthiols). If the amount
of ink solution is limited only a drop can be placed on top of the structured stamp surface. After a given time excess solution is sucked off using a pipette or blown away
with a stream of nitrogen. When using this kind of loading procedure impurities or
possible precipitated material from the ink solution can adsorb to the stamp surface and
disturb the ink transfer during printing. A third method called contact inking uses an
ink pad which could be either a flat slab of PDMS that was soaked with ink or a glass
slide coated with a thin layer of ink solution. Utilizing such an ink pad only the raised
stamp regions/features are wetted and broadening of the printed patterns by vapor diffusion from the recessed non contacting regions is almost prevented. Since PDMS is
hydrophobic it is supposed that polar inks remain at the surface of the stamp whereas
unpolar molecules diffuse into the bulk. To this end when working with unpolar inks
the stamp can be rinsed with pure solvent after wetting in order to remove possible contaminations. The wettability of the stamp surface with polar inks and the uniformity of
printed patterns can be improved by oxidizing the PDMS surface using a oxygen plasma.
Nevertheless, independent on the technique used for wetting and the nature of the ink
molecules after wetting the stamp has to be dried thoroughly to evaporate excess solvent
and avoid pattern distortions due to a possible swelling of the stamp.
After drying the loaded stamp is gently pressed onto the substrate surface typically for
5-15 seconds. Thereby the ink molecules are transferred to the surface only at those
regions where the relief surface of the stamp contacts it. In this regions the molecules
assemble into more or less ordered structures. The ordering upon printing as well as the
transfer onto the substrate are not fully understood processes depending on a variety of
parameters like concentration and properties of the ink molecules. A scheme of stamp
fabrication and printing is presented in Figure 3.2.
The total pattern area that can be transferred depends on the outer dimensions of the
template pattern. Typically areas between 0.25 and 100 cm2 can be printed at once with
edge resolution better than 50 nm. The lateral dimensions of the printed features can
23
Figure 3.2.: Fabrication process of a PDMS stamp and preparation of a lateral structured
SAM by microcontact printing
typically range from 50 nm to several cm. However, patterns with lateral dimensions
in the nanometer region are difficult to obtain. Lateral diffusion of the ink results in
broadening of the printed pattern and low mechanical stability of the elastomeric stamp
results in distortions of the stamp pattern e.g. collapse and deformation (Fig. 3.4)). In
order to prevent deformations of the pattern the aspect ratio of height to width (h/w) or
feature size to separation distance (w/d) should be between 0.2-2 or 0.1-0.5, respectively
[51, 49]. In addition to this fabrication limitations distortions of the printed pattern can
be caused by applying a nonuniform pressure during printing or swelling of the stamp
material (a number of nonpolar organic solvents cause swelling of PDMS). Drying the
stamp after loading with ink prevents swelling. Diffusion caused pattern deformation
due to rest solvent in-between the relief structures is also reduced. To overcome these
technical limitations other elastomers [52, 53] or composite stamps (e.g. PDMS supported by glass [54] or hard PDMS [55]) have been proposed.
24
Figure 3.3.: Further treatment of laterally structured SAMs. A) filling remaining gold
areas with a second thiol. B) fabrication of a gold-silicon microstructured substrate
After printing the patterned substrate can be used in different ways. When immersing
the patterned substrate in a solution containing another adsorbate with different chemical/physical properties the bare gold regions are filled with those molecules (Fig. 3.3 A).
Applications for patterned SAMs with different properties are manifold. For example in
cell biology, cell interaction with different surfaces can be studied or cell adhesion and
growth can be controlled. Thereby size of the pattern and its molecular composition
25
Figure 3.4.: Different types of stamp deformations
can affect the DNA synthesis, cell growth and protein secretion of the attached cells.
The ability to pattern defined arrays of immobilized cells makes the construction of new
types of whole-cell-based sensors possible.
Even the ability of SAMs to act as an etch resist and protect the underlying gold form
dissolving can be used to generate microstructured gold-silicon substrates (Fig. 3.3 B).
Exposure of a patterned SAM (e.g. ODT) to an etching solution results in the dissolution of gold at those regions not protected by the SAM. The resulting microstructures of
gold can again be used as masks, which protect the underlying silicon from alkaline etch
so that only the unprotected regions are dissolved. The shape of the etched features is
determined by the etching conditions.
In comparison to other structuring techniques µCP provides some advantages. The technique is experimentally simple and can be performed in ambient chemical lab conditions
without access to clean rooms or photolithographic equipment. Furthermore a single
master can be used for the production of many stamps and even each stamp can be used
many times. In contrast to electron-beam or scanning probe lithography pattern transfer
occurs simultaneously over a large area, allowing a high speed of transfer. Shortcoming
is the difficulty to fabricate a pattern with more than two different functionalities. This
would require precise alignment of multiple printing processes.
3.2. Scanning probe lithography (SPL)
With the invention of AFM and STM researchers noticed that the surfaces under investigation can be altered under certain conditions. To this end scanning probe lithography techniques were developed which provide controlled modification of surfaces at the
nanoscale. In the last decade different types of SPM lithography were introduced, for
instance:
• Bias-induced nanolithography [56]: Writing is accomplished by applying pulses or
elevated bias voltage between a conductive tip and a conductive or semiconductive
26
3.2.1. Nanografting
surface coated with an insulating SAM. The surface under the tip becomes oxidized
under elevated bias providing a reactive site for attaching new molecules. Often
nanopatterns written with bias-induced nanolithography do not produce height
changes so that no topographic contrast is observed. However differences in lateral
force (friction) are recognized due to the different material properties.
• Dip-pen nanolithography (DPN) [57]: AFM tips are used as pens that are coated
with a molecular ink for writing on clean substrates in air. Writing mechanism
involves the transfer of molecules to the surface through a nanoscopic meniscus that
forms between the tip and the substrate. Therefore humidity conditions have to be
stringently controlled. After writing an uncoated tip can be used to characterize
the fabricated structures.
• Catalytic probe lithography [58]: a tip coated with catalytic agent that reacts with
the terminal groups of a SAM is used for writing. As the tip touches areas of the
surface the molecular coating of the tip induces a catalytic reaction and chemically
changes the terminal groups of the SAM.
• Nanografting which is described in detail in the next section.
Precisely engineered nanostructures obtained by scanning probe lithography allow the
exploration of chemical and biochemical reactions under spatially well-defined and controlled environment. Studies on those nanostructures provide fundamental information
on tip-surface interactions, structures, and properties on a nanoscopic level and serve as
a useful guide in the development of nanoelectronic devices, biosensors, and biochips.
When choosing a suitable SPL method researchers need to consider what types of surface
functionalities are desired, the imaging medium, and the nature of the surfaces under
investigation. For further and more detailed information on scanning probe lithography
the reader is referred to published reviews [59, 60, 61].
3.2.1. Nanografting
Nanografting is a SPM-based lithography technique that was first reported in 1997 by
Gang-yu Liu and coworkers [62]. It allows lateral structuring of molecules within a matrix SAM with nanometer resolution. A review on nanografting was recently published
[63].
Contact mode imaging of SAMs or other soft matter is generally performed with a minimal force applied to the tip (< 1 nN). The low pressure exerted by the tip can still
result in small local deformations and a very small shear force during scanning that is
too low to damage the surface. Increasing the local pressure increases the deformation
and the tip penetrates into the SAM. At a specific pressure the shear force becomes so
high that it causes a cleavage of the gold-thiolate bond since this is the weakest bond
in the organic molecule (binding energies for Au-S, S-C, C-C, C-H are 30, 171, 145, 81
27
kcal/mol, respectively). Consequently, the adsorbate molecule dissolves from the surface.
Fabricating structures using nanografting includes three steps (Fig. 3.5). The first step
is to image the SAM surface with low loading force in a liquid medium containing a
different kind of adsorbate and select an area for nanografting. In the second step the
selected area is scanned with a loading force higher than the displacement threshold.
When the loading force on the tip is increased chemisorbed molecules within the SAM are
removed. Immediately after the displacement adsorbate molecules from the surrounding
solution chemisorb onto the freshly exposed surface following the scanning track of the
tip. Finally the fabricated structure is characterized at a low imaging force.
Figure 3.5.: Nanografting process in three steps 1) the surface is characterized under a
low loading force 2) zooming in a target area and displacing the molecules by increasing
the applied force 3) a larger area is scanned to characterize the fabricated structure
under a low loading force
If nanografting is performed in a pure solvent that does not contain possible adsorbates
the process is called nanoshaving. In this case molecules within the SAM are removed
leaving a hole in the SAM. There is only a narrow range of force over which the tip can
selectively desorb the thiols. Too high loading forces on the tip during the displacement
process will cause plastic deformation of the underlying gold substrate. If the loading
force is too low, adsorbed molecules will not be displaced. The displacement threshold
force depends on the local structure of the SAM, the geometry of the AFM tip (tip’s
curvature radius), and the fabrication environment (e.g. roughness of the gold surface).
Therefore, the appropriate force for nanografting must be determined in situ for each
experiment. Next to the force also the solvent influences the desorption procedure. In
solvents with little solubility for the displaced molecules most of them remain weakly
attached to the gold substrate or the SAM interface. In those solvents the displacement is mostly reversible and nanografting is less successful [64]. Better solvents allow
the desorbed molecules to dissolve and diffuse away from the fabrication site more easily.
Formation of a high quality SAM requires many hours. In contrast, a high quality
nanografted structure can be created in seconds. This implies that adsorption is accelerated upon nanografting. While natural self-assembly from solution occurs in two steps
(striped phase and crystalline phase, see section 2.3.2.2 (page 11) the striped phase was
28
3.2.1. Nanografting
not observed in nanografting. Unlike the case of natural (unconstrained) growth freshly
exposed areas of gold produced in nanografting are spatial confined by the AFM tip
and the surrounding SAM at the boundary of the scanning track. Because the adsorbates do not have sufficient room to assemble in a lying-down orientation they adsorb
onto the gold with a favored standing-up configuration directly [62, 65]. Therefore the
assembling process in nanografting is called spatially confined self-assembly (SCSA).
Even if natural growth and nanografting show different adsorption pathways it was
demonstrated
that the structure of nanografted octadecanethiol (ODT) show the same
√ √
◦
( 3x 3)R30 overlayer as an ODT SAM but the nanografted structure contains less
defect sites [62, 65]. Experiments with different thiols show that SCSA induces a closepacking of the molecules within the fabricated patterns [66, 67].
Nanografting can produce a great variety of patterns and shapes with a broad range of
functional groups. The lateral resolution depends on the geometry and sharpness of the
tip apex. The smallest feature yet produced by nanografting is a 2 x 4 nm2 dot of ∼32
thiol molecules [68]. Changes and/or modifications of the fabricated patterns can be carried out very quickly and easily without repeating the entire fabrication procedure [66].
By successive change of the nanografting solution (solution containing the desired adsorbate) it is also possible to produce multiple patterns with different functional groups
by successive fabrication steps. However, a challenging task is to overcome the relatively
low fabrication speed which is too slow for mass production because of the serial patterning process. Typical writing speeds for a single tip are about 2 µm/s. A parallel
fabrication of nanostructures could be obtained using one or two-dimensional AFM tip
arrays. Prototype arrays of 55000 AFM probes have recently been developed by Mirkin
and co-workers and were applied in DPN [69]. Another critical point is the changing tip
geometry induced by friction and wear during nanografting which influences the writing
process and the quality of the fabricated structures.
3.2.1.1. Nanografting of biological molecules
One of the main advantages of nanografting with respect to biological applications is that
the sample remains immersed in liquid throughout the whole experiment (nanopatterning, rinsing, characterization). Especially in case of protein immobilization this plays an
important role since those generally denature during drying and lose their functionality.
Different studies on direct immobilization of proteins onto gold surfaces were published
by Scoles and coworkers [70, 71, 72, 73]. Since SAMs offer a multitude of functional
terminal groups even immobilization of proteins on nanografted structures were studied
particularly with regard to the development of surface protein assays at the nanoscale
[74, 75, 68, 76, 77].
Furthermore nanografting has been applied to characterize and control the orientation
and packing of DNA on surfaces. In contrast to unconstrained self-assembly spatially
confined self-assembly results in case of DNA in almost fully stretched conformations of
the DNA molecules [45, 78, 79]. Different factors impact the molecular packing density
of DNA during nanografting:
29
• line density parameter [78, 79]:
The line density parameter (S/A) describes how often a given area is grafted. It
depends on the tip radius at the point of contact with the surface (R), the number
of scan lines (N) and the length of the nanografted structure (L): S/A = R·N
. For
L
a S/A ≤1 the nanografted lines do not overlap whereas for a line density parameter
of 2 each line is grafted twice. Very recently Mirmomtaz et al. demonstrated that
especially at line densities between 1 and 15 the packing density is highly sensitive
to the chosen line density [79].
• nanografting force [78]:
Low applied loading forces during nanografting creates low density nanostructures
because the tip is not able to remove all SAM molecules. To this end only fewer
DNA molecules can chemisorb onto the freshly exposed gold.
• concentration of the nanografted adsorbate [78]:
DNA packing density is highly dependent on thiolated ss-DNA concentration in
solution. Low concentrations result in a reduced packing density.
Very recently Mirmomtaz et al. [79] reported on the ability of nanografting to systematical control DNA surface coverage. Furthermore they demonstrated hybridization of
very high density nanostructures and proof that molecular density is not the decisive
factor for inhibited hybridization in high density DNA SAMs. The authors attribute
this result to an improved ordering (less entanglement) of the DNA molecules within
the fabricated nanostructures due to SCSA. In addition they were able to quantify hybridization efficiency of DNA nanostructures by studying its compressibility at different
loading forces.
30
4. Theory of scanning probe
microscopy (SPM)
Contents
4.1. Atomic force microscopy (AFM) . . . . . . . . . . . . . . . .
32
4.1.1. AFM set-up and operation modes . . . . . . . . . . . . . . .
32
4.1.1.1. Contact mode . . . . . . . . . . . . . . . . . . . . .
32
4.1.1.2. Intermittent mode (”Tapping mode” or Dynamic mode) 35
4.1.1.3. Non-contact mode . . . . . . . . . . . . . . . . . . .
36
4.1.1.4. Other scanning modes/techniques: . . . . . . . . . .
36
4.2. Chemical force microscopy (CFM) . . . . . . . . . . . . . . .
37
31
Chapter 4. Theory of scanning probe microscopy (SPM)
In 1981 Gerd Binning and Heinrich Rohrer invented the first type of SPM which was
termed scanning tunneling microscope (STM). For their discovery they were awarded
the Nobel Prize for Physics in 1986. The STM records the variation in tunneling current
between a sharpened tip and a sample as the tip passes over the atomically corrugated
surface. As tip-sample distance decreases and increases due to height features, the tunneling current increases and decreases respectively obeying an exponential relationship.
By keeping the tunneling current constant the tip-surface distance can be precisely maintained. However, the restriction to electrically conductive samples limited the choice of
materials to be investigated.
Today the term scanning probe microscopy (SPM) covers a variety of methods which
enable the observation and manipulation at the atomic scale. All methods have in common that a miniature cantilever with a sharp tip probes the surface. The interaction
between the surface and the tip differ among the different modes of operation resulting in different contrast mechanism. Among them mechanical, electrostatic, optical, or
magnetic information. Nevertheless, still today the primary use of SPM is to monitor
the 3D surface topography.
4.1. Atomic force microscopy (AFM)
In 1986 Gerd Binnig and coworkers invented the atomic force microscope (AFM) which
allows imaging insulating surfaces as well as electrically conductive surfaces. Like the
STM, the AFM also uses a very sharp tip to probe surface morphology. In the AFM,
van der Waals forces acting between the tip and surface provide the contrast mechanism
instead of the tunneling current. Thereby the usability of SPM methods was dramatically
enhanced because then the resolution is only dependent on the sharpness of the tip.
The information offered by AFM goes beyond physical dimensions (e.g. topography)
of a surface. For example, in intermittent mode the local hardness, the elasticity, and
the electrostatic properties of a surface can be visualized. Furthermore the intermittent
mode allows to observe soft materials without damaging the surface. Therefore it is now
an established and often used tool in biology and life sciences.
4.1.1. AFM set-up and operation modes
There are three main imaging modes in AFM based on different types of forces: contact
mode, intermittent mode (tapping mode), non-contact mode. In the following these
modes will be outlined. The general set-up and operation of an AFM is introduced with
the following description of the contact mode.
4.1.1.1. Contact mode
As the name already implies in contact mode the tip is continually in contact with the
surface and ”feels” the surface topography. In this mode short-range repulsive forces
(e.g. van der Waals forces) act between tip and surface (Fig. 4.1, light green). When
32
Figure 4.1.: Force as function of probe-sample separation and corresponding AFM operation modes
imaging in air the tip contacts the surface through the adsorbed water layer on the
sample surface.
A sharp tip attached to a micro-cantilever is raster scanned across the sample to monitor
the surface topography. Depending on the AFM design either the tip can be moved or
the sample. In order to accomplish three-dimensional sub-Ångstrom movements piezoelectric scanners are typically used in SPM. Scanning the tip is favored for large, heavy
samples. When moving the sample it is most common to use a piezoelectric tube scanner with the sample mounted on top. The flexible cantilever is used as a spring to
measure the force between the tip and the sample. Depending on the local interaction
between tip and sample the cantilever bends toward (attractive interactions) or away
from the surface (repulsive interactions). A laser focused on the back of the cantilever
and deflected onto a position-sensitive photodiode allows monitoring movements of the
tip. Most AFMs use a 4-segment photodiode where top-down difference signal is indicative for the interaction force between tip and sample. As the surface is scanned the tip
follows the surface topography and moves according to the surface profile up and down.
The changes in vertical deflection are proportional to the forces between tip and scanned
33
Figure 4.2.: AFM laser beam deflection system. A laser beam is focused on back of the
cantilever and deflected onto a 4-segment photodiode. The top-bottom signal difference
gives information on the normal force. Left-right deflections provide information about
lateral forces. The lower drawing depicts the feedback loop.
34
surface. More precisely the tip surface interaction force can be calculated using Hooke’s
law F = −k∆x, where F is the interaction force, k is the stiffness of the lever and z is the
deflection of the cantilever due to the applied force. In order to follow the topography
with a constant force applied to the surface an electronic feedback loop is necessary. The
feedback loop maintains the position of the deflected laser spot constant by applying a
correction voltage to the z portion of the piezo scanner. From the voltage used to drive
the piezo z-scanner the required height adjustments are known. Recording the height
information point-by-point yields a topography image. In praxis the SPM controlling
software often allows the user to define a ”setpoint” parameter. This value tells the
feedback loop which vertical difference signal to maintain during scanning. Thus the
user defined setpoint is proportional to the imaging force (loading force applied by the
tip).
Noise is a critical factor in AFM operation. In order to achieve highest resolution the
microscope must be isolated from noise in its surroundings, such as floor vibrations,
acoustic, electrical and/or optical noise source.
A shortcoming of contact mode operation is that the tip can damage and/or deform
soft samples. These effects are in praxis minimized by applying as low loading forces
as possible and by selecting appropriate tips. Because the tip is in hard contact with
the surface, the stiffness of the lever needs to be less than the effective spring constant
holding atoms together, which is on the order of 1-10 nN/nm. Most contact mode levers
have a spring constant of < 1 N/m. A softer cantilever means that in comparison to a
hard cantilever lower loading forces are required to give the same vertical deflection signal
onto the photodiode. Often low loading forces are desired when imaging in contact mode.
However, the downside of soft cantilevers is the risk of picking up noise more easily.
Advantageous is that contact mode operation allows fast scanning. Furthermore this
mode can be performed in ambient and liquid environment. With regard to ”hard”
samples it allows very high resolution up to the atomic level (e.g. atomic resolution of
inorganic crystals).
4.1.1.2. Intermittent mode (”Tapping mode” or Dynamic mode)
This mode belongs to the family of AC modes, which refers to the use of an oscillating
cantilever. A stiff cantilever is oscillated at its resonance frequency in the attractive force
regime (Fig. 4.1, light red). The lowest points of oscillation extends into the repulsive
force regime, so that the tip intermittently touches or ”taps” the surface. When the tip is
close to the surface the free oscillation is damped. By maintaining a constant oscillation
amplitude a constant tip-sample interaction is achieved. In this case the setpoint value
reflects the deflection amplitude of the oscillation on the position-sensitive photodiode.
A high setpoint value means less damping of the oscillation resulting in a larger tipsample distance and hence a lower imaging force.
Intermittent mode allows high resolution images of samples that are easily damaged
and/or loosely adsorbed to a surface. Therefore it is the scanning mode of choice for
softer matter samples (e.g. polymers, biological samples). Since the tip contacts the
35
surface only very shortly during scanning frictional forces which exert lateral forces onto
the sample are almost eliminated. A shortcoming of the intermittent mode is the much
slower scan speed in comparison to contact mode. In addition intermittent mode in
liquids is a challenging task due to the strong damping by the liquid.
Typically very stiff cantilevers having resonance frequencies of 200-400 kHz, and spring
constants of more than 10 N/m are employed in intermittent mode. In intermittent
mode the resolution becomes better with higher resonance frequencies. Therefore the
cantilevers need to be stiffer. High forces are present between tip and surface in the very
short time interval when the tip taps onto the surface. Despite of this intensive contact
most organic surface are not harmed by the high normal force which has been attributed
to the visco-elastic effect. Often for operation of the intermittent contact mode in liquid
soft ”contact mode” cantilever are used. Capillary and meniscus forces between surface
and tip are vanishing but the liquid is strongly damping the vibration of the cantilever.
Not only the oscillation amplitude but also the resonance frequency is strongly reduced.
4.1.1.3. Non-contact mode
Also this operation mode belongs to the family of AC modes. As in intermittent mode a
stiff cantilever is oscillating in the attractive regime, but in contrast to the intermittent
mode the sample surface is not touched by the tip during oscillation (Fig. 4.1, light blue).
The oscillation amplitude is reduced allowing stable non-contact imaging by probing
weak attractive van der Waals forces. Nevertheless the non-contact mode is less stable
than contact or dynamic mode since the forces between tip and surface are much smaller
than in the other modes. Consequently upon perturbations there is a higher possibility
that the non-contact regime is lost e.g. the tip jumps out of contact or the tip jumps into
full contact. Due to capillary forces the non-contact mode is difficult to control under
ambient conditions. Very stiff cantilevers are needed so that the attraction does not
overcome the spring constant of the cantilever, but the lack of physical contact between
tip and sample means that this mode causes the least disruption. Advantageous is that
the forces between tip and sample are quite low, on the order of pN. A shortcoming is
that usually ultra high vacuum is needed to obtain good images. As discussed above
a contaminant layer (e.g. water) on the sample surface can easily impair the measured
signal and interfere with the tip-sample interaction.
4.1.1.4. Other scanning modes/techniques:
Lateral Force Microscopy (LFM):
Beyond up and down movements of the cantilever also left and right deflections of the
laser spot on the four-segment photodiode can be monitored. Physically the left and
right movements are interpreted as torsion of the cantilever due to frictional forces acting on the tip. Lateral deflection can give information about the friction between the
tip and the sample, and can show areas that may have the same height, but different
chemical properties. In order to maximize lateral forces the fast scan direction must be
perpendicular to the long axis of the cantilever. Typically the lateral force is recorded
36
4.2. Chemical force microscopy (CFM)
simultaneously to topographic information in contact mode.
Phase imaging:
In Phase mode imaging, the phase shift of the oscillating cantilever relative to the driving
signal is measured. This phase shift can be correlated with specific material properties
(e.g. friction, adhesion, and visco-elasticity) that affect the tip-sample interaction. The
phase information is usually recorded simultaneously with topography in intermittent
mode.
Constant height mode:
In this mode the same line is scanned twice. First the topographic information is recorded
using contact or intermittent mode. Subsequently the topographic information is used
to track the tip at a constant distance above the surface. While maintaining a constant
height different properties such as magnetic or electric forces can be probed.
Force modulation:
The tip (or sample) is oscillated at a high frequency while maintaining in the repulsive
force regime. This means the tip does not leave the surface during the oscillation cycle.
Upon every up and down of the tip during oscillation the slope of the force-distance
curve is measured which is correlated to the sample’s elasticity. In this way relative
stiffness of surface features is probed and mapped.
4.2. Chemical force microscopy (CFM)
Although AFM enables characterizing nano-mechanical properties of the sample with
resolutions down to the atomic scale a drawback is the lack of chemical specificity.
To overcome this inherent limitation in 1994 Lieber and coworkers developed a AFM
technique based on chemically functionalized tips which provide chemical specificity [80].
By utilizing tips covered with a well-defined layer of molecules, this techniques can be
used to (i) probe intermolecular forces between different chemical groups (one is bound
to the tip and the other kind covers the surface), (ii) measure surface energetics on a
nanometer scale, (iii) determine pK values of the surface acid and base groups locally,
and (iv) map the spatial distribution of specific functional groups and their ionization
state using lateral force imaging. The development of various techniques to chemically
modify an AFM tip has enormous expanded the use of this technique. A review on CFM
is given by Noy et al. [81].
When comparing CFM with SNIM some issues have to be considered:
(i) CFM can probe only the topmost layer of sample whereas SNIM has a specific
penetration depth of several tens of nanometers allowing subsurface imaging.
(ii) because in CFM the probe must be functionalized a priory knowledge of the sample
composition is necessary
37
(iii) in CFM the chemical interaction has to be separated from other interactions like
capillary forces, morphology, or mechanical properties.
38
5. Theory of infrared (IR) spectroscopy
Contents
5.1. Infrared spectroscopy . . . . . . . . . . . . . . . . . . . . . . .
40
5.2. Fourier transform infrared (FTIR) spectroscopy . . . . . . .
44
5.2.1. Infrared reflection absorption spectroscopy (IRRAS) . . . . .
46
5.3. IR microscopy or microspectroscopy . . . . . . . . . . . . . .
48
39
Chapter 5. Theory of infrared (IR) spectroscopy
In infrared spectroscopy, IR radiation is passed through a sample. Some of the radiation
is absorbed by the sample and some of it is transmitted. The resulting infrared spectrum
contains absorption peaks which correspond to the frequencies of vibrations between
the bonds of the atoms the material consists of. Therefore, infrared spectroscopy is
known as vibrational spectroscopy. Since each material is a unique combination of
atoms and no two molecular structures produce exactly the same infrared spectrum, it
represents a molecular fingerprint of the sample. Positions and intensities of vibrational
bands can be used to confirm or identify the presence of a particular group, whereas
spectral correlations can be used to access structural and environmental information on
selected groups. IR spectroscopy is particulary non-intrusive, requires small amount of
sample and can be easily coupled with microscopy providing spatial resolution in order
to produce chemical maps of samples. Now-a-days infrared spectroscopy is a widely used
analytical technique.
5.1. Infrared spectroscopy
Infrared refers to the region from 800 nm to 1000 µm in the electromagnetic spectrum
between the visible and microwave range. Commonly this spectral region is separated
into three parts:
near-IR (NIR)
800 nm < λ < 2.5µm
12500 < ν̃ < 4000cm−1
excitation of vibrational overtone and combination bands
mid-IR (MIR)
2.5 < λ < 50µm
4000 < ν̃ < 200cm−1
excitation of fundamental vibrational modes
far-IR (FIR)
50 < λ < 1000µm
200 < ν̃ < 10cm−1
excitation of scaffold oscillation and rotational modes
Infrared radiation can be absorbed by molecules and excites molecular vibrations and
rotations within the molecule. In contrast to vibrational transitions, the rotational transitions are only visible in gas phase spectra. Most molecules have characteristic infrared
active vibrational modes whose vibrational energy corresponds to the energy of electromagnetic waves in the MIR region. Thereby absorptions bands above 1500 cm−1 can
be assigned to single functional groups whereas absorption bands below 1500 cm−1 are
characteristic for the molecule itself. To this end the MIR region is often called ”fingerprint region”.
40
In infrared spectroscopy it is common to use wavenumbers ν̃, in units of cm−1 , rather
than the wavelength λ since the wavenumber is directly proportional to energy
ν̃ [cm−1 ] = 104 /λ [µm] =
E
hc
(5.1)
ν̃ = wavenumber, λ = wavelength, E = energy, h = Planck’s constant, c = speed of light
Excitation of molecular vibrations and the formation of an infrared spectrum can be
described by a simple classical mechanical model. Two atoms are approximated by the
model of a harmonic oscillator composed of two masses m1 and m2 joined by a spring
with a force constant k at the equilibrium distance r0 between the masses (Fig. 5.1).
If the distance between the masses is expanded or compressed, as during a vibration,
a restoring force Fback arises and the system vibrates around its equilibrium distance.
According to Hooke’s law this force is proportional to the displacement:
Fback = −k∆r
(5.2)
Fback = withdrawing force, ∆r = r-r0 = displacement, k = force constant
Figure 5.1.: Scheme of a vibrating diatomic molecule and potential curves of a harmonic
(A) and anharmonic (B) oscillator
41
The frequency of the vibration is given by:
s
m1 · m2
1
k
with µ =
ν0 =
m1 + m2
2π µ
(5.3)
ν0 = vibrational frequency, k = force constant, µ = reduced mass, m = mass
According to this formula a higher vibrational frequency results from a higher force
constant k, i.e. a stronger bond, and/or smaller masses. Examples for the influence of
bonding strength are given in Table 5.1.
In a classical harmonic oscillator the potential energy Epot is
1
Epot = k∆r2
2
(5.4)
Fback = restoring force, ∆r = r-r0 = displacement, k = force constant
and rises during displacement parabolic around the equilibrium distance. Solving the
Schrödinger equation shows that the vibrational energy is quantized:
1
1
Ev = hν0 (v + ) = k∆r2
2
2
(5.5)
E = energy, h = Planck’s constant, v = vibrational quantum number, ν0 = vibrational frequency,
k = force constant, ∆r = r-r0 = displacement
The lowest energy level is E0 = 21 hν0 and the next level will be E1 = 32 hν0 .
Consequently the energy states are equidistant. According to the selection rule only
transitions to the next energy level are permitted: ∆v = ±1. The absorbed energy
difference needs to be
∆E = Ev+1 − Ev = hν0 .
(5.6)
E = energy, ν0 = resonance frequency
The model of the harmonic oscillator is a very simplified approximation for a two atomic
molecule and is not able to explain some phenomena such as molecular dissociation. A
better description is the model of an anharmonic oscillator or the so-called Morse potential. According to this model large vibrational amplitude could result in a dissociation
force constant [105 dyne/cm]
single bond:
5
double bond:
10
triple bond:
15
examples
C−C, C−O, C−N
C=C, C=O, C=N
C≡C, C≡N
vibrational frequency [cm−1 ]
800 < ν̃ < 1300
1500 < ν̃ < 1900
2000 < ν̃ < 2300
Table 5.1.: Correlation between bond strength and vibrational frequency
42
of the molecule and moreover the potential curve is much steeper during bonding compression due to the Pauli principle.
Solving the Schrödinger equation with the Morse potential shows the vibrational states
are not equidistant anymore. Instead their differences in energy decrease in direction
of the dissociation energy D. According to the selection rule not only transitions with
∆v = ±1 are possible but even higher transitions with ∆v = ±2, 3, ..., n , so-called
vibrational overtones. Their transition probability and therefore their intensity decreases
strongly with increasing ∆v. Generally only the ground state E0 = hν20 is populated at
room temperature. Furthermore appearance and intensity of vibrational bands depends
on the transition dipole moment of the vibrating groups in the molecule. Absorption
and emission of radiation can only occur if the transition dipole moment changes during
vibration
∂~µel
6= 0
(5.7)
∂r
µ
~ = dipole moment, r = displacement
Consequently all vibrations symmetric to a symmetry center of a molecule are IR inactive (i.e. forbidden) because the dipole moment does not change. A molecule consisting
of N atoms has a total of 3 N degrees of freedom, corresponding to the cartesian coordinates. Three of these degrees are translational movements in x-, y-, and z-direction,
three are rotational degrees around the three main moments of inertia, and the remaining correspond to fundamental vibrations. It follows:
degrees of freedom for a linear molecule:
degrees of freedom for a nonlinear molecule:
n = 3N − 5
n = 3N − 6
Molecular vibrations can be divided into two groups (Fig. 5.2):
(i) stretching ν : displacement oriented parallel to the bonding axis. The bond distance
changes periodically. If two atomical groups are symmetrically equivalent and
their vibrations are coupled they are called symmetric νs or antisymmetric νas
stretching vibrations depending if the atoms vibrate into the same direction or
not, respectively.
(ii) deformation δ : the bonding angle changes periodically and the bond distance remains almost constant. Here, there are also symmetric and antisymmetric vibrations.
A summary of typical vibrational bands of biomolecules is given in the appendix in
Table B (page 184).
43
Figure 5.2.: Typical molecular vibration modes
5.2. Fourier transform infrared (FTIR) spectroscopy
The source of radiation in a spectrometer generally emits a broad range of frequencies.
In order to monitor the variation of absorption with frequency conventional infrared
spectrometer contain a dispersive element which separates the emitted frequencies of
the radiation source so that only a nearly monochromatic beam passes the sample at a
time. Those dispersing elements are generally prisms or gratings which separate radiation with different frequencies into different spatial directions. The light intensity at
each frequency transmitted by the sample is recorded.
As the name implies FTIR spectrometer use a fourier transform technique for spectral
detection and analysis. The heart of such a spectrometer is a Michelson interferometer,
a device for analyzing the frequencies present in a composite signal. A Michelson interferometer employs a beam splitter which divides the beam coming from the sample into
two beams. One beam reflects off a flat mirror which is fixed in a plane. The other beam
reflects off a flat mirror whose distance to the beam splitter varies continuously within a
certain range (typically a few millimeters). Thus, a varying path difference ∆x between
both beams is introduced. When the two beam components recombine at the back of
the beam splitter there is a wavelength dependent phase difference between them and
they interfere either constructively or destructively depending on the difference in the
path lengths. The detected signal oscillates as the two components alternately come
into and out of phase as the path difference is changed. The resulting signal is called
an interferogram. If the radiation has wavenumber ν̃ the intensity of the detected signal
due to radiation in the range of wavenumbers ν̃ to ν̃ + dν̃ varies with x as:
I(x, ν̃) dν̃ = I(ν̃) (1 + cos 2πν̃x) dν̃
(5.8)
I (x, ν̃) = detected signal intensity, ν̃ = wavenumber, x = path difference
Hence, the interferometer converts the presence of a particular wavenumber component
in the signal into a variation in intensity of the radiation reaching the detector. An
actual signal consists of radiation containing a large number of frequencies and the total
intensity at the detector I(x) is the sum of contributions from all the wavenumbers
present in the signal:
44
5.2. Fourier transform infrared (FTIR) spectroscopy
Figure 5.3.: A) Dispersive spectrometer B) FTIR spectrometer with a Michelson interferometer
Z
I(x) =
∞
Z
I(x, ν̃) dν̃ =
0
∞
I(ν̃) (1 + cos 2πν̃x) dν̃
(5.9)
0
I(x) = detected signal intensity, ν̃ = wavenumber, x = path difference
Thus, the interferogram contains the intensities of all infrared frequencies ”encoded”. In
order to ”decode” the absorption spectrum of the sample from the recorded values I(x)
a mathematical fourier transformation is performed giving I(ν̃, the variation of intensity
with wavenumber. Specifically:
Z ∞
I(ν̃) = 4
{I(x) − 0.5I(0)} cos(2πν̃x) dx
(5.10)
0
I(ν̃) = intensity, I(x) = detected signal intensity, I(0) = I(x) with x = 0, ν̃ = wavenumber,
x = path difference
The resulting spectrum is identical to that from a conventional dispersive spectrometer.
A major advantage of the Fourier transform procedure is that all frequencies emitted by
the source are monitored simultaneously resulting in an increased measurement speed
(seconds instead of minutes). Moreover even the signal-to-noise ratio increased for two
reasons:
45
1) the amount of light reaching the detector at any given time is higher than in a
conventional instrument. All light available from the source is continually delivered
to the detector in a FTIR spectrometer while the slits in a conventional instrument
restrict the amount of light.
2) The random noise is reduced by signal averaging. Since only a few seconds are
needed to record an interferogram many can be acquired in the time needed to
complete a conventional spectrum with a dispersive spectrometer. Random noise
is reduced by the square root of the number of spectra averaged together.
In order to avoid instrumental characteristics, such as influences from the emission spectrum the radiation source or from components guiding the beam a so-called reference
or background spectrum is recorded. This is normally a measurement without a sample
(I0 (ν)). Subsequently (or simultaneously), the sample spectrum (I(ν)) with identical
configuration of the spectrometer is recorded. For receiving the ”pure” sample absorption
spectrum the logarithmic ratio of reference and sample spectrum is calculated according
to Lambert-Beer’s Law:
I(ν)
A(ν) = −log10
(5.11)
I0 (ν)
A(ν) = absorbance spectrum, I0 (ν) = reference spectrum, I(ν) = sample spectrum
5.2.1. Infrared reflection absorption spectroscopy (IRRAS)
The increased signal-to-noise ratio of FT instruments allows infrared spectroscopy of
very small amounts of species such as organic thin films. For the characterization of
SAMs FT-IRRAS is a very powerful tool.
A configuration commonly used to analyze infrared spectra of thin organic films is grazing angle reflection geometry. A piece of metal with a thin film on top is positioned so
that the incoming light is reflected under a large angle of incidence (> 80◦ relative to the
surface normal). As the light passes through the thin film, a small amount is absorbed
by the film via vibrational excitation. In order to maximize the absorbance the angle
needs to be very large.
Reflection spectra can differ from those obtained by transmission methods. The electric
field in close proximity to a metal surface generates mirror dipoles since the electrons inside the metal are easily polarized. In dependence of the orientation of the polarization
of the light different phenomena occur. S-polarized light (electric field perpendicular
to the plane of incidence) generates an antiparallel oriented mirror dipole in the metal
surface which almost cancels out with the incoming s-polarized light. P-polarized light
(electric field parallel to the plane of incidence) causes a mirror dipole that adds to the
incoming p-polarized light. Consequently electric field components parallel to the surface vanish, while field components perpendicular to it are enhanced. This gives rise to
what is called the ”surface” selection rule. The general selection rule for absorption of IR
light requires a vibration producing a change in the transition dipole moment. Upon adsorption of the molecules to a surface the molecular mobility is strongly limited. Due to
46
Figure 5.4.: Signal processing in a FTIR spectrometer. Signal intensity is recorded as
function of the mirror displacement (interferogram). Mathematical fourier transformation gives the intensity as function of the wavelength (transmission spectrum). The
logarithmic ratio of reference and sample spectrum yields the sample absorption spectrum. (Spectra are recorded from a 6-mercapto-1-hexanol SAM (sample) and a bare
gold substrate (reference).)
the frozen molecular orientations all vibrating dipole components parallel to the surface
will cancel out due to the surface selection rule. All vibrational modes having transition
dipole moments normal to the metal surface are enhanced in external reflection infrared
spectroscopy. When mixing the molecules in KBr pellets volume transmission spectra
can be recorded. In these spectra all IR active vibrations show up. Thus comparison of
the transmission and reflectance spectra provides information about molecular orientation of the adsorbed molecules. In the same way molecules in disordered films will be
more similar to their transmission spectra. This is because disordered molecules retain
a higher degree of flexibility and thus the vibrational signal will be an average over the
different molecular orientations. Consequently the peak width will broaden. In addition,
47
Figure 5.5.: Schematic representation of the surface selection rule. S-polarized light
(perpendicular to the plane of incidence) cancels out at a metal surface with a thin film
on top. P-polarized light (electric field parallel to the plane of incidence) is enhanced.
Consequently only vibrational modes having transition dipole moments normal to the
surface are detected in IRRAS.
the effects of the surface selection rule become less pronounced with increasing distance
from the surface. Gases or liquids above the thin film on the metal surface are not
affected by the surface selection rule so that both polarization components are equally
absorbed.
A very important issue in IRRAS is the reference sample. When investigating SAMs
on gold the easiest available reference is a bare gold substrate. But one has to be
aware that gold is instantaneously covered with a physisorbed layer of water, hydrocarbons, and other organic compounds from the atmosphere under ambient laboratory
conditions ([19] and references therein) which can effect the vibrational bands of the
sample. For instance physisorbed hydrocarbons on the gold surface show an absorption
in the methylene region around 3000 cm−1 and can cause a disappearance or negative
methylene peaks in the sample spectrum. Better references are hydrophobic SAMs which
prevent adsorption of contaminants onto the surface. Very well-suited are perdeuterated
alkylthiolates since they are hydrophobic, form densely packed SAMs and the deuteration avoids spectroscopical overlap with the C-H vibrations.
A technique that requires no additional reference is polarization modulated infrared
reflection spectroscopy (PM-IRRAS). This technique measures differences in the absorption of p- and s-polarized light simultaneously. The p-polarized light contains vibrational information from bulk and surface adsorbed molecules while the s-polarized
light is sensitive to the molecules in the bulk only. When applied towards SAMs on gold
a big advantage of this technique is that no additional reference spectrum needs to be
recorded. This enhances spectral accuracy, avoids errors due to contaminated reference
surfaces and speeds up the measurement process.
5.3. IR microscopy or microspectroscopy
Mapping the chemical state of a specimen is of great importance in a variety of scientific
fields. Infrared microscopy has developed into a powerful analytical tool. It provides
spatially resolved chemical contrast, enables in situ measurements, is non-intrusive and
requires only small amounts of sample. Typically, a conventional microscope working
at IR wavelengths is combined with a fourier-transform infrared (FTIR) spectrometer
48
5.3. IR microscopy or microspectroscopy
to acquire a spectrum at each pixel of the image. A shortcoming is the limitation in
spatial resolution given by the Abbe diffraction limit: a lens-based imaging instrument
cannot achieve a better spatial resolution than approximately half of the wavelength
(about λ/2). However, the spatial resolution achieved in IR microspectroscopy is approximately 2λ due to throughput deficiencies and optical aberrations. Present available
FTIR microscopes achieve a resolution ranging from a few microns to a few tens of microns depending on the wavelength.
Another method of IR microspectroscopy is Raman scanning microscopy. In this case,
a similar but complementary type of molecular information (molecular vibrations) as in
IR microscopy is revealed, while the spatial resolution is defined by the spatial profile
of the focused laser beam, which has a wavelength in the visible or near-IR. Spatial
resolution close to 1 µm is achievable. The Raman effect is a nonlinear process with
a much smaller cross-section (∼10−28 to 10−30 cm2 ) than a typical single-photon electronic absorption cross-section (∼10−18 to 10−20 cm2 ). As a consequence Raman spectra
show a dramatically reduced intensity in comparison to IR absorption spectra (roughly
a factor of 10−10 ). In order to obtain a good signal-to-noise ratio Raman microscopy
experiments require either high power pump beams that bear the risk of thermal damage
to the sample or resonance effects like in surface enhanced raman scattering (SERS),
since otherwise integration times will be impractically long.
49
6. Theory of scanning near-field
optical microscopy (SNOM)
Contents
6.1. Historical development of SNOM . . . . . . . . . . . . . . . .
53
6.2. SNOM configurations . . . . . . . . . . . . . . . . . . . . . . .
55
6.2.1. Aperture SNOM . . . . . . . . . . . . . . . . . . . . . . . . .
56
6.2.2. Apertureless or scattering SNOM (s-SNOM) . . . . . . . . .
59
6.3. Theory of s-SNOM . . . . . . . . . . . . . . . . . . . . . . . .
60
6.3.1. Near- and far-field of a Hertzian dipole . . . . . . . . . . . . .
60
6.3.2. Quasi-electrostatic theory . . . . . . . . . . . . . . . . . . . .
61
6.3.2.1. Tip Apex . . . . . . . . . . . . . . . . . . . . . . . .
62
6.3.2.2. Tip-sample interaction - Dipole mirror-dipole interaction/model . . . . . . . . . . . . . . . . . . . . . .
64
6.3.3. Background suppression . . . . . . . . . . . . . . . . . . . . .
67
6.3.3.1. Modulation of the tip-sample distance (direct detection scheme) and higher harmonics detection . . . .
68
6.3.3.2. Homodyne interferometric detection . . . . . . . . .
69
6.3.3.3. Heterodyne interferometric detection . . . . . . . .
70
6.3.4. Applications of SNOM . . . . . . . . . . . . . . . . . . . . . .
72
6.3.4.1. Aperture SNOM in the optical regime . . . . . . . .
72
6.3.4.2. Scattering SNOM in the optical regime . . . . . . .
73
6.3.4.3. Aperture SNIM . . . . . . . . . . . . . . . . . . . .
73
6.3.4.4. Scattering SNIM . . . . . . . . . . . . . . . . . . . .
73
6.3.4.5. s-SNOM in the THz regime . . . . . . . . . . . . . .
74
51
Chapter 6. Theory of scanning near-field optical microscopy (SNOM)
An everlasting dream of human beings is to understand nature in its smallest details.
Since the development of lenses and their use to magnify objects much effort was spend
to improve lateral resolution. After the invention of compound microscopes consisting
of two or more lenses further development was devoted to optimization of lens shape
and quality, correction of aberrations and fabrication of sophisticated combinations of
illumination and collection lenses.
In the 1870’s Ernst Abbe and Lord Rayleigh demonstrated that lateral resolution in
optical systems is limited by diffraction to 1 :
∆x ≥ 0.61
λ
λ
= 0.61
n sin α
NA
(6.1)
∆x = lateral resolution (distance between two objects to be resolved);
λ = wavelength of the incident light; n = refractive index;
sin α = half angle through which the light is collected; NA = numerical aperture
With regard to this limitation the easiest way to improve lateral resolution is to reduce
the wavelength of light. Unfortunately, with smaller wavelengths the photon energy
increases. Damage due to high energy photons strongly limits the usage of UV-light
in life sciences (e.g. live cell imaging). Due to wave-particle-duality even electron and
neutron microscopes exist. With x-ray, electron and neutron microscopes the energy
induced damage becomes even worse. Therefore, these techniques are generally classified
as destructive imaging techniques. The resolution of these techniques is determined by
their short wavelengths. However, even at those small wavelengths the law for the
diffraction limit is still fulfilled. Since the resolution of light microscopes reaches nearly
the diffraction limited resolution, aberrations of the optical components in electron- and
neutron microscopes drastically limit the achievable resolution in practise. Table 6.1
outlines the resolution and (dis-)advantages of some popular microscopy techniques.
Another further improvement in optical microscopy was the invention
√ of confocal microscopy. With this kind of microscope the resolution improved by ∼ 2 [83] and since
it suppresses out-of-focus fluorescence the signal-to-noise is additionally improved. In
the last two decades several new optical imaging techniques have been developed to
beat the diffraction limit such as 4Pi-microscopy [84] or stimulated-emission-depletion
microscopy [85]. But all of them are based on the use of fluorophores. Attaching bright
fluorophores in an oriented quantitative manner to any biological species is difficult and
always bears the risk of loosing biological functionality.
A fundamentally different approach for obtaining a high lateral resolution is SPM (outlined in chapter 4 (page 31)). In contrast to classical microscopy SPM requires no sample
illumination for visualization. Instead sample properties (e.g. topography) are probed
with a very small tip (nano finger). Therefore it contains in comparison to optical
microscopy complementary information.
1
Abbe’s law in its original form did not contain the factor 0.61 (Equation 6.1). This factor comes from
the Rayleigh criterion and is based on diffraction at a circular aperture.
52
6.1. Historical development of SNOM
Technique
optical microscopy
confocal LSM
(MMM)-4Pi
SEM
environmental SEM
(ESEM)
Typical
λ
[nm]
400-700
(VIS)
Best lateral
resolution
[nm]
∼250
400-700
(VIS)
∆x:∼200
∆z:∼1000
400-700
(VIS)
2.5-12 pm∗
(electrons)
∆x:∼100
∆z:∼500
0.4 [82]
∼2
Advantages and shortcomings
+ many different contrast mechanisms
+ easy to handle
+ imaging in liquids
+ imaging in liquids
- fluorescence dyes
- slow due to scanning
- fluorescence dyes
+ spatial image (high depth of field)
- conductive and metallic samples
- vacuum
+ gas chamber (up to 20 torr)
+ uncoated insulating and wet samples
- expensive
Table 6.1.: Achievable resolution and (dis-)advantages of popular microscopy techniques; ∗ 2.5 pm at 200 kV and 12.3 pm at 10 kV, LSM = laser scanning microscopy,
MMM = multifocal, multiphoton microscopy, SEM = scanning electron microscopy
Scanning near-field optical microscopy is a concept that detects simultaneously optical
and topographical surface information (a combination of ”nano eye” and ”nano finger”).
It improves optical resolution by circumventing the use of lenses and consequently breaks
Abbe’s diffraction-based resolution limit. In contrast to far-field lens-based microscopy,
where the illumination source as well as the detection system is placed in large distances
from the sample ( λ), in near-field microscopy one of both (either the illuminating
or detection system) is positioned in close proximity to the sample ( λ). The very
small distance between probe and sample allows to probe the near-field (non-propagating
waves) of the sample which contain information about the localized electric charge distribution.
A review about high resolution microscopy techniques was published by Garini et al.
[83].
6.1. Historical development of SNOM
The first concept to circumvent diffraction limited resolution was proposed 1928 by Edward H. Synge [86]. He suggested to illuminate a sample through an aperture that is
significantly smaller than the employed wavelength of light while keeping the sample-toaperture separation at a distance much smaller than the wavelength of light - that is in
53
Figure 6.1.: Synge’s proposed concept to overcome the diffraction limit (1928). Light
passes through a sub-wavelength aperture in an opaque screen and illuminates a sample
that is placed within the near-field ( λ) of the aperture.
the range of the optical near-field. For image formation the sample is scanned point-bypoint. Unfortunately, the idea was ahead of his time and technically not realizable, due
to a lack of powerful light sources, like lasers, precise distance control, and fabrication
processes for tiny apertures. In the course of time different publications on theory of optical fields behind small apertures appeared e.g. from Bethe in 1944 [87], Bouwkamp in
1950 [88, 89], O’Keefe in 1956 [90] and McCutchen in 1967 [91]. Nevertheless it was until
1972 when Ash and Nicholls demonstrated the first ”super-resolution aperture scanning
microscope” [92] and achieved a lateral resolution of λ/60 at a frequency close to 10 GHz
(λ = 3 cm). An important invention for the further development of near-field microscopy
was the scanning tunneling microscope by Binnig and Rohrer in 1982 [93] which allows
to approach and scan a sharp tip in close proximity to a sample surface. Already two
years later, in 1984, the first scanning near-field microscope working with visible light
was reported by Pohl et al. [94]. They succeeded in fabricating tiny apertures at the end
of a metal coated probe. With those they were able to recognize details of 25 nm size
using light with a wavelength of 488 nm resulting in a resolving power of ∼ λ/20. Another independent work published in the same year by Lewis et al. reported also about
fabrication and characterization of tiny apertures [95] and proposed a similar scheme for
a near-field microscope. Not until 1991 when Betzig and coworkers [96] demonstrated
SNOM probes made from a single-mode optical fibre and shear-force-distance detection
research in the field of near-field optic developed drastically. Especially the possibility
to gain highly resolved optical information simultaneously with topographic information is interesting for many applications. In 1985 Wessel proposed a new concept for
near-field microscopy based on apertureless probes [97] allowing a further improvement
of lateral resolution in comparison to aperture-based near-field microscopy, especially
at long wavelengths like infrared [98, 99], THz [100] or microwaves [101]. The significance of this long wavelength imaging is that new types of material contrast become
for the first time accessible to sub-micrometer microscopy, namely contrast due to resonant vibrational absorption enabling to image and identify chemical composition of a
nanostructured material.
54
6.2. SNOM configurations
6.2. SNOM configurations
Scanning near-field microscopy can be classified into two main families: (i) aperture
SNOM and (ii) scattering or apertureless SNOM (Fig. 6.2). In both types of SNOM a
very small probe, which can be an emitter, collector, or scatterer, is brought in close
proximity to the sample. If the distance is smaller than the wavelength the probe can
interact with the near-field of the sample which contains smallest information of the
surface charge distribution (≤ λ/2). The probe is scanned in a raster pattern over the
surface of the sample. Because of the exponential decay of the near-field the probe has to
be kept at a constant distance from the sample during scanning. Otherwise the intensity
changes caused by different probe-sample distances will shadow the desired contrast
mechanism. The distance between probe and sample is usually controlled through a
feedback mechanism, which can be either:
• normal force feedback, i.e. the standard feedback mode of an AFM, which is
limited to bent optical fibres and AFM cantilevers with holes
• shear force or tuning fork feedback, for which a straight fiber is mounted to
a tuning fork or a piezo, which excites the tip to oscillate parallel to the sample
surface. During approach the resonance frequency of the fibre is detuned/damped
with respect to the oscillation of the driver due to the action of shear forces leading
to a decrease in amplitude and a shift in phase. When reaching a pre-defined setpoint value, approach is stopped and the value is maintained by the feedback
system. Advantageous is that this kind of feedback control mechanism is nonoptical.
Figure 6.2.: Different types of near-field scanning optical microscopes
55
As the probe is scanned over the sample, the signal intensity is recorded point by point.
SNOM instruments are able to record simultaneously topography and near-field signal.
This is extremely valuable since both data sets can be compared to determine correlations
between the physical structure and optical contrast.
6.2.1. Aperture SNOM
Aperture SNOM is based on the concept proposed by Synge in 1928 [86] (Fig. 6.1).
Light propagates through a sub-wavelength optical aperture and illuminates the sample
or detects light from it. Most aperture SNOMs employ either a tapered optical fibre
with a metal coating at which only the foremost end is left uncoated or AFM cantilevers
with a hole in the center of a pyramidal tip.
The most important properties concerning aperture probes are the aperture diameter,
which determines the spot size and therefore the lateral resolution, and the light intensity at the aperture, which should be sufficient to obtain a good signal-to-noise ratio.
Both properties are closely correlated to each other. Only a tiny fraction of light - typically 10−3 to 10−6 - coupled into an aperture probe is emitted through its aperture.
The rest of the light is either reflected back into the waveguide or absorbed in the metal
coating, leading to a considerable heating of the metal. The three main factors that affect transmission efficiency of aperture probes are: the taper cone angle, the size of the
aperture, and the damaging threshold of the metal coating. Below a critical diameter
defined as
dc = 0.6λ/n
(6.2)
dc = cut-off diameter, λ = wavelength, n = index of refraction
light propagation in a circular waveguide becomes evanescent and its intensity decays exponential in propagation direction. The amount of light that reaches the probe aperture
depends on the distance separating the aperture and the so called cut-off diameter dc .
Figure 6.3.: Scheme of light propagating in SNOM probes with different cone angles.
Dashed lines show the impact of the cone angle on topographic contrast
56
6.2.1. Aperture SNOM
This distance is smaller in tapers with large cone angles than in those with smaller ones
[102] (Fig. 6.3). A serious drawback is that with increasing taper cone angle in fact
the transmission greatly improves but at the same time the suitability for simultaneous
topographic imaging with high resolution decreases (Fig. 6.3). Nevertheless operation
below the cut-off diameter imposes severe power losses (output power of some nW in the
visible) and prevents a sub-micrometer resolution for longer wavelengths such as infrared
since their cut-off diameter is larger than in the visible. Furthermore transmitted light
intensity at a sub-wavelength aperture depends strongly on the aperture diameter. The
transmission coefficient decreases drastically with decreasing aperture size [87, 88]:
4
d
T ∝
λ
(6.3)
T = transmission coefficient, d = diameter of aperture, λ = wavelength
Therefore aperture size is limited. Typical aperture diameters are 50-100 nm in the
visible range and around 1 µm in the infrared region. Attempting to overcome the small
transmission by using high input power is limited to a certain threshold power. Higher
input power causes thermal damage of the metal coating at the tapered end of the probe.
For this reason aperture SNOMs in the visible are typically operated with input power of
≤ 10 mW. However, Hong et al. demonstrated that measurements in liquid environment
allow the use of much higher laser powers (up to 2 W in the infrared region) [103].
In aperture SNOM different operation modes can be applied whereby the probe can
either serve as emitter or collector. In collection mode the sample is externally illuminated, e.g. through a focused beam. This illumination generates evanescent waves at
the sample surface which are converted by the aperture into propagating waves inside
the fiber and conducted through the fiber to a suitable detector. In illumination mode
the light is coupled into an aperture probe and guided to its tapered end, towards the
aperture. The evanescent field at the aperture interacts with the fine structure of the
sample leading to a conversion into a propagating wave that can be detected in the farfield. Moreover SNOM can be performed in transmission or reflection. The most applied
modes are displayed in Figure 6.4. Choice of operation mode depends primarily on the
application. For example opaque samples are not suited for transmission mode whereas
when working with fluorophores collection mode is unfavorable because the sample is
externally illuminated over a large area which leads to excessive photobleaching within
the entire illuminated zone around the tip, even before it can be scanned.
Aperture SNOM is presently the most popular SNOM technique. All commercial instruments currently available employ the aperture SNOM technique. In most cases
conventional optical microscopes are extended by a SNOM zooming stage. Further and
more detailed information on aperture-based SNOM is given in a review by Hecht et al.
[104].
57
Figure 6.4.: Aperture SNOM operation modes: A) Illumination in transmission, B)
Collection in transmission, C) Collection in TIR illumination (photon tunneling), D)
Collection in oblique reflection, E) Illumination in oblique reflection, F) Illumination
and collection in reflection
Operating aperture SNOM in the infrared and THz region is still a challenging task with
respect to high quality IR fiber fabrication. Commonly used silica optical fibers exhibit
strong absorptions in the mid-IR region and are therefore unsuitable in this spectral
range. Unfortunately most IR transmitting glasses have a limited transmission range
in the mid-IR and/or undesirable mechanical properties. Widely used materials for infrared fibers are chalcogenide glasses which are transparent in the wavelength region
from 2-12 µm. They are based on the chalcogen elements S, Se, and Te. Adding other
elements such as Ge, As, and Sb leads to the formation of stable glasses. The best lateral resolution achieved with such fibers was 100 nm using a free electron laser as light
source [105, 106, 107]. Another material for IR-fibers is silver halide (AgClBr) [108, 109].
58
6.2.2. Apertureless or scattering SNOM (s-SNOM)
6.2.2. Apertureless or scattering SNOM (s-SNOM)
Commonly this type of near-field working with a sharp tip instead of an aperture is called
apertureless SNOM or ASNOM although it is more precise to call it scattering SNOM
(s-SNOM). In s-SNOM a sharp tip with a sub-wavelength diameter at its apex scans a
sample in close proximity as in a STM or AFM. In addition, the tip is illuminated by a
focused beam of an external radiation source. Since the tip is externally illuminated it
is preferable to scan the sample and hold the tip fixed. The light scattered from the tip
is recorded in the far-field whereby input and output directions could be freely chosen.
Backscattering configuration has the advantage that only one focusing element has to
be adjusted. Different configurations of s-SNOM are presented in Figure 6.5. A main
advantage of s-SNOM is that standard AFM tips can be used as probes. In s-SNOM
the tip has two optical functions: (i) to intercept and reradiate incoming light like an
optical antenna and (ii) to supply a concentrated longitudinally polarized optical field
at the tip apex to probe the optical response of a nearby sample material. Briefly,
when exposed to an electromagnetic wave the tip acts as an antenna and generates a
locally concentrated near-field at its apex. This near-field is modified by the presence
of a sample. As a consequence of this near-field interaction, even the scattered light
from the tip’s apex detected in the far-field is modified and contains information on
Figure 6.5.: s-SNOM operation modes: A) illumination and collection in reflection, B)
evanescent excitation mode, C) illumination and collection in transmission, D) illumination in transmission, collection in reflection
59
the local dielectric properties of the sample. A detailed description of the theory of
apertureless SNOM follows. Spatial resolution in s-SNOM depends mainly on the tip’s
curvature radius which is typically about 10 to 40 nm, and therefore results in much
better resolution than aperture SNOM. The best resolution attained so far with an
apertureless microscope was 1 nm by Zenhausern et al. in 1995 [110]. In comparison
to aperture SNOM the scattering-type can be operated at different wavelengths (e.g.
VIS and IR) without much effort when changing the wavelength. Routinely used probe
materials are PtIr and gold (Au) due to their high polarizability. Even silicon nitride
(Si3 N4 ) and silicon tips can provide a sufficient scattering signal [111]. In case of silicon
tips the usual covering SiO2 layer needs to be removed by etching.
6.3. Theory of s-SNOM
The terms near- and far-field describe regions with different physical properties around
any electromagnetic radiation source. The boundary between these two regions is not
distinct and only vaguely defined: the near-field is roughly speaking the region within
a radius r λ while the far-field is the region for which r λ. When describing the
electromagnetic field of the source in the Fourier space it consists of a broad spectrum
of spatial frequencies. High spatial frequency Fourier components are only present near
the source and decay exponentially, radially along the object normal. This region is
called near-field zone. For low spatial frequencies the waves propagate in z-direction
towards the observation plane decaying by 1/r. These Fourier components represent
the far-field. With regard to the diffraction limit lenses placed in the far-field can only
collect spatial frequencies smaller than NA/λ. However, SNOM is able to measure the
rapidly decaying near-field (surface-bound) components of a radiating object by placing a
small probe within the near-field region of the object. The electric near-field of an object
contains detailed information about the object structure and its charge distribution that
are not available in the far-field.
6.3.1. Near- and far-field of a Hertzian dipole
The terms near- and far-field will be explained using the emitted field of the easiest
point dipole, the Hertzian dipole.
A hertzian dipole consists of two oscillating opposite point charges. Its dipole moment
is:
p~ = p~0 · eiωt
(6.4)
p = dipole moment, ω = angular frequency, t = time
The emitted electric and magnetic field components in spherical coordinates are:
60
6.3.2. Quasi-electrostatic theory
2
p
ṗ
Er =
+
cos θ
4πε0 r3 cr2
p
ṗ
p̈
1
+
+
sin θ
Eθ =
4πε0 r3 cr2 c2 r
ṗ
p̈
1
Hφ =
+
sin θ
4π r2 cr
1
2p
1ω
=
+i 2
cos θ
4πε0 r3
r c
p
1
1 ω 1 ω2
=
+i 2 −
sin θ
4πε0 r3
r c
r c2
p
1
1 ω2
=
iω 2 −
sin θ
4π
r
r c
(6.5)
(6.6)
(6.7)
Er = radial component of electric field, Eθ = angular component of the electric field,
Hφ = magnetic field component, p = dipole moment, r = distance with respect to the dipole center,
c = speed of light, ṗ, p̈ = temporal derivatives of the dipole moment, ω = angular frequency,
ε0 = dielectric constant
For large distances (r c/ω = λ/2π) the 1/r terms dominate the electric field and
the radial component (Er ) cancels out. Consequently transversal electromagnetic waves
form with their electric and magnetic field components perpendicular to each other.
Those waves propagate only perpendicular to the dipole moment because their intensity
is highest at the equator (θ = π/2) and cancels out at the poles (θ = 0, π). This zone
~ 2 decreases with 1/r2 is called far-field.
where the detectable intensity |E|
For small distances (r < c/ω = λ/2π) the 1/r3 term becomes dominant and the inten~ 2 drops rapidly with 1/r6 . Because the electric field components do not reach the
sity |E|
far-field they are referred as non-propagating or non-radiating waves. The zone around
the dipole in which these waves decay is called near-field. Within these zone even components parallel to the dipole moment are emitted so that the near-field decays radially
around the dipole. In contrast to the far-field the near-field contains information about
the spatial charge distribution of the dipole.
The interaction energy between oscillating electric point dipoles (atoms, molecules, or
mesoscopic particles) is proportional to 1/r6 if these are in near-field contact. If the
dipoles are located in the far-field zone of each other the interaction energy is proportional to 1/r2 . When the near-field interaction dominates light retardation effects often
can be neglected, and quasistatic approaches are adopted in the solution of specific problems. Solids can be imagined to be composed of an assembly of point dipoles arranged in
a regular lattice. Summing up all electric field contributions from the various regularly
spaced dipoles results in an exponentially decaying evanescent field.
In scattering near-field microscopy only light scattered from the very apex of the tip
contains near-field information. Because the tip is typically much smaller than the
wavelength the probe’s apex can be approximated as a point or Rayleigh scatterer.
Probe-sample interaction can be qualitatively described by quasi-electrostatic theory. In
case in which the sample can be approximated as a small object such as a small particle
(e.g. gold nanoparticle), probe and sample are both approximated as polarizable spheres
(dipole-dipole interaction). In order to describe the optical interaction of a probe with
61
a planar sample like a SAM the slightly modified dipole mirror-dipole theory [112, 113]
can be employed. The latter theory is even applicable for the samples investigated in
this thesis. Therefore only the dipole-mirror dipole theory will be explained in detail.
6.3.2.1. Tip Apex
The probe’s apex can be approximated as a polarizable sphere because only the light
scattered from the very apex of the tip contains near-field information (Fig. 6.6). When
applying an electric field E0 the sphere becomes polarized with dipole moment
p = αE0
(6.8)
p = dipole moment, α = polarizability, E0 = electric field
The polarizability α describes the tendency of distorting the normal charge distribution
upon applying an external electric field. According to electrostatic theory polarizability
of a small spherical particle (radius a wavelength λ) is:
εprobe − εmedium
3
(6.9)
α = 4πa
εprobe + 2εmedium
α = polarizability, a = radius of curvature, ε = complex dielectric function
In order to calculate the scattering and absorption cross-sections of the probe, Mie theory
can be applied which describes scattering from spherical particles. In case of scattering
on particles much smaller than the wavelength of light Mie theory reduces to Rayleigh
Figure 6.6.: In quasi-electrostatic theory the tip apex is approximated as polarizable
sphere with radius a. The tip apex acts as a Mie scatterer when illuminated with an
external electric field.
62
approximation (Rayleigh scattering). Scattering depends on the optical properties of the
material. According to the Maxwell equations refractive index and dielectric constant
are related to each other:
√
n = εµ
(6.10)
n = refractive index, ε = dielectric constant, µ = permeability
For dielectric materials the relative permeability is µ ≈ 0 following:
ε = n̂2
ε0 + iε00 = n2 − κ2 + 2niκ
ε0 = n2 − κ2
(6.11)
ε00 = 2nκ
and
(6.12)
ε = complex dielectric constant, n̂ = complex refractive index
The scattered electric field is:
Esca = σE0 = seiφ E0
(6.13)
Esca = scattered electric field, E0 = incident electric field, σ = scattering cross-section,
s = scattering amplitude, φ = scattering phase
Approximating the tip furthermore as small sphere (radius a λ or ka1, k = 2π/λ)
in a medium the scattering cross-section σ is:
σ = σsca + σabs
(6.14)
with
σsca
2
8π 4 6 nˆr 2 − 1 =
k a 2
3
nˆr + 2 σsca
with nˆr =
n̂probe
n̂medium
ε
2
probe
−
1
8π 4 6 εmedium
=
k a 3
ε εprobe + 2 medium
(6.15)
(6.16)
σsca = total scattering cross-section, k = wave vector, a = radius,
nˆr = relative complex refractive index, n̂ = complex refractive index, ε = complex dielectric constant
and for materials with κ 6= 0 light is not only scattered but can also be absorbed. The
absorption cross-section is given by:
σabs
2
nˆr − 1 2
= 4πka 2
nˆr + 2 3
with nˆr =
n̂probe
nmedium
ˆ
(6.17)
63
σabs
 ε

 ε probe − 1 
medium
= 4πka3 Im  εprobe + 2 
(6.18)
εmedium
σsca = total scattering cross-section, k = wave vector, a = radius,
nˆr = relative complex refractive index, n̂ = complex refractive index, ε = complex dielectric constant
Rearranging Equation 6.9 and insertion in Equations 6.16 and 6.18 results in:
σsca =
k4 2
|α|
6π
(6.19)
σsca = scattering cross-section, k = wave vector, α = polarizability
σabs = kIm{α}
(6.20)
σabs = absorption cross-section, k = wave vector, α = polarizability
6.3.2.2. Tip-sample interaction - Dipole mirror-dipole interaction/model
In theory the sample is modeled as a half space whose interface is located at z = r − a
below the tip with z λ. Each sample material is characterized by its dielectric constant εs and its polarizability. Due to the sample’s polarizability the close-by induced
probe dipole can influence the charge distribution in the sample under the probe. The
locally distorted charge distribution can be described by an image dipole located at a
distance of 2r from the probe dipole. Depending on the polarization of the external
~ 0 probe and image dipole can enhance or cancel each other out.
electric field E
When E0 is polarized parallel to the surface the electric field of the probe dipole
Eprobe (r) = −
pprobe
4πr3
(6.21)
E = electric field, p = dipole moment, r = distance from dipole center to sample surface
induces an image dipole oscillating 180◦ out of phase with the tip dipole. Due to the
opposite orientations of probe and image dipole the resulting total dipole moment is
strongly reduced or even cancels out completely (Fig. 6.8 B). Therefore a perpendicular
polarization was used during experimentation. In the following description of near-field
theory only the perpendicular polarization will be considered. When E0 is polarized
perpendicular to the surface the electric field of the probe dipole
Eprobe (r) =
pprobe
2πr3
E = electric field, p = dipole moment, r = distance from dipole center to sample surface
64
(6.22)
Figure 6.7.: Scheme of a dipole-mirror dipole interaction. The tip apex is approximated
as polarizable sphere. When exposed to an electric field a mirror dipole in a closeby sample is induced. As consequence of the near-field interaction between both the
scattered light from the tip apex is modified and contains information on the local
dielectric properties of the sample.
induces an image dipole p’:
0
p = βpprobe =
εsample − 1
εsample + 1
pprobe
(6.23)
p’= dipole moment of mirror dipole, β = constant of proportionality, p = dipole moment,
ε = dielectric function
β is a constant of proportionality relating the polarizability of the sample to the polarizability of the probe αprobe :
εsample − 1
αsample = βαprobe =
αprobe
(6.24)
εsample + 1
α = polarizability, β = constant of proportionality, ε = dielectric function
Probe dipole pprobe and image dipole p’ are pointing in the same direction resulting in
an enhanced total dipole moment (Fig. 6.8 A).
The total electric field acting at the apex of the tip is composed of the coherent sum
of E0 and the contribution of the dipoles. This leads to the following modified probe
dipole moment:
65
Figure 6.8.: Orientation of external electric field components A) Electric field components perpendicular to the sample surface and parallel to the tip are enhanced B)
Electric field components parallel to sample surface and perpendicular to the tip are
surface cancel out.
p0
= αprobe E0 +
2π(2r)3
β
= αprobe E0 +
pprobe
16πr3
αprobe
E0
=
αprobe β
1 − 16πr
3
pprobe
(6.25)
p = dipole moment, α = polarizability, E0 = external electric field,
r = distance from dipole center to sample surface, β = constant of proportionality
The effective polarizability of the coupled probe and mirror dipole system can be described by the following equation:
ef f
α⊥
=
=
αprobe + αprobe β
α
β
probe
1 − 16πr
3
αprobe + αprobe β
1−
αprobe β
16π(z+a)3
αef f = effective polarizability, α = polarizability, β = constant of proportionality,
r = distance from dipole center to sample surface, a = tip curvature radius, z = r-a
66
(6.26)
6.3.3. Background suppression
In order to calculate the scattering and absorption cross-sections of the probe when
interacting with a sample the polarizability α in Equation 6.19 and 6.20 have to be
substituted by the effective polarizability αef f
σsca =
k4
|αef f |2
6π
(6.27)
σsca = scattering cross-section, k = wave vector, α = polarizability
σabs = kIm{αef f }
(6.28)
σabs = absorption cross-section, k = wave vector, α = polarizability
The electric field of the light scattered from the apex of the tip is proportional to αef f .
The detected intensity of the scattered light is proportional to the square of its electric
field.
Remark: It has to be noted that the relative simple dipole-mirror dipole theory is not
able to describe properly the near-field interaction on multilayer samples. Depending on
the thickness of the upper layer of a multilayer sample and the penetration depth of the
tip’s near-field not only the first layer is probed by the tip but also the underlying one.
Such a sample is for instance a SAM on gold. The thickness of a SAM is typically less
than 10 nm and therefore less than the probed depth of the tip. To this end also the
dielectric response from the gold layer has to be considered since it effects the probe’s
polarizability.
A multilayer theory was recently proposed by Moiseev et al. [114] and Aizpurua et al.
[115] and will be considered in future theoretical predictions on s-SNIM applied to SAMs
on gold.
One of the main challenges in scattering SNOM is to extract the light coming from the
tip’s apex over a large background signal. Illumination of the scattering probe is usually
achieved through a free-space focusing element, like a lens, a parabolic mirror, or an
objective, either in reflection or transmission configuration. In any case the illuminated
area is much larger than the tip’s apex including the shaft and cantilever of the tip and
the sample surface. As demonstrated in the previous chapter only the light scattered
from the apex of the tip is influenced by near-field interactions with the sample surface.
Light scattered from any other element sums up to a large background signal |EB |2 . In
order to detect the weak scattered light from the apex of the tip |EN |2 efficient methods
for background suppression need to be implemented. Therefore in the following different
methods for background suppression will be discussed.
67
Figure 6.9.: The area illuminated by the focused laser beam is much larger than the
tip’s apex. Consequently light scattered from other elements (e.g. sample, tip’s shaft,
cantilever) sums up to a large background signal.
6.3.3.1. Modulation of the tip-sample distance (direct detection scheme) and
higher harmonics detection
The most convenient way to extract the near-field information containing signal was
already demonstrated in the first realization of near-field optical microscopy and is based
on a modulation of the probe-sample distance [92]. Oscillating the tip height results in
a sinusoidal modulation of the scattered light from the tip. Using a lock-in amplifier
each non-modulated signal is suppressed (e.g. stray light from the sample). Since all
scattered signals are generated from the same driving field they add coherently. Thus
the signal detected is a complicated sum of background and near-field components and
an interference product of both 2 :
∗
Idetector = (EB + EN )(EB∗ + EN
)
= |EB |2 + 2|EB ||EN | cos(φB − φN ) + |EN |2
(6.29)
I = intensity reaching the detector, EB = electric field background,
EN = electric field containing near-field information, E ∗ = complex field conjugated to E, φ = phase
Generally the background signal is much larger than the near-field signal EB EN so
that |EB |2 dominates the detected intensity. The background signal can be composed
of unmodulated portion of light (e.g. stray light) and a modulated portion of light (e.g.
a standing wave forms via interference effects, see below). When modulating the tip at
2
68
EN = σE0 and EB = σB E0 with σ = σsca + σabs
a frequency f the unmodulated background light is discarded by usage of a lock-in amplifier. Please note that the modulated background light does not disappear and is still
collected by the lock-in amplifier. Upon approach of the sample surface remaining background signal and near-field information containing signal show a different behaviour.
Since the distance dependence of the near-field signal is strongly non-linear (see Equation 6.26) a sinusoidal modulation at frequency f of the tip-sample distance generates
higher-harmonic components of the scattering signal:
σ = σ0 + σ1 cos(f t) + σ2 cos(2f t) + ... + σn cos(nf t)
(6.30)
σ = scattering cross-section, f = modulation frequency, n = 1, 2, 3, ...
At the same time the background scattering varies much more smoothly on approach
thus containing weaker harmonics. However within the distance λ from the surface also
interference effects between light scattered from the tip’s shaft and the sample surface
have to be considered (Fig. 10.8). The interfering waves produce a standing-wave which
introduces some higher harmonics [116] which are however less pronounced than the
higher harmonics of the near-field containing signal.
σ b = σ0b + σ1b cos(f t) + σ2b cos(2f t) + ... + σnb cos(nf t)
(6.31)
σ = scattering cross-section, f = modulation frequency, n = 1, 2, 3, ...
Nevertheless the different scales of z-dependence of the near-field influenced signal and
the background signal allow improving the signal/background ratio EN /EB according
to (λ/2πa)n with higher harmonics n [113, 117]. With respect to Equation 6.29 demodulation at higher harmonics results in a signal were the interference term between
background and near-field |EB ||EN | cos(φB − φN ) rises over the pure background term
|EB |2 .
Because the background signal cannot be fully suppressed due to the coherent superposition of the different scattered fields, detection at higher harmonics is not sufficient
to extract a pure near-field signal. Nevertheless this detection scheme is experimentally
much simpler than those described in the following. And although the contrast interpretation is difficult the interference term might be beneficial for contrast formation
since the background field EB can amplify a very weak near-field signal EN [118, 119].
In order to overcome interpretation difficulties due to background effects and gain pure
near-field signal higher harmonics detection was combined with interferometric detection
[117, 120].
6.3.3.2. Homodyne interferometric detection
In homodyne interferometric detection [120] a Michelson-type interferometer is implemented to a higher harmonic detection scheme (Fig. 6.10 A). In this configuration the
movable mirror of the interferometer is switched between two positions with a distance
of λ/8 (corresponding to a phase difference of 90◦ ). At the first position a maximum
69
output signal proportional to sn cos(φn ) is detected (n is the demodulation order). This
signal is stored and the mirror is moved to the second position. In this case the reference
beam is 90◦ phase shifted and the signal is proportional to sn sin(φn ). Finally from the
two detected signals sn and φn can be calculated. In contrast to sole higher harmonics detection an additional reference field has to be considered resulting in a far-field
detected intensity
∗
)
Idetector = (EB + ER + EN )(EB∗ + ER∗ + EN
2
2
2
= |EB | + |ER | +|EN | + 2|EB ||EN | cos(φB − φN )
|
{z
}
filtered out by lock-in
(6.32)
+ 2|EB ||ER | cos(φR − φB ) +2|EN ||ER | cos(φR − φN )
|
{z
}
EN = electric field containing near-field information, ER = electric field reference,
E ∗ = complex field conjugated to E, φ = phase
The intensity after the lock-in detection is thus:
I = |EN |2 +
2|E ||E | cos(φ − φN )
| B N {z B
}
+2|EN ||ER | cos(φR − φN )
(6.33)
suppressed by higher harmonic detection
E ∗ = complex field conjugated to E, φ = phase
According to the equations above all non-modulated terms can be extracted by lock-in
detection. Unfortunately this is not the case for the term resulting from the interference between background field and near-field because also this term is modulated by the
tip frequency. As described before it can be suppressed by lock-in detection at higher
harmonics (explanation see above). But if the reference beam from the interferometer
is strong enough its interference with the near-field signal exceeds the background-nearfield interference manifold. Even when strongly suppressed the effect of the undesired
interference term between background field and near-field cannot be completely eliminated or controlled using homodyne interferometric detection.
6.3.3.3. Heterodyne interferometric detection
Heterodyne detection was employed for the first time in 2000 by Hillenbrand et al. [117].
It enables to map the optical near-field amplitude and phase and to separate the pure
near-field response from all background signals allowing therefore even nanoscale optical
mapping of topography-rich objects. In contrast to homodyne interferometry where the
interference occurs between a reference beam (local oscillator) and a probing beam at the
same wavelength (or carrier frequency) in heterodyne interferometry prior to detection
the reference beam is frequency shifted by ∆ω (beat frequency) relative to the probing
beam. For instance frequency shift can be obtained using an optical acoustic modulator
70
Figure 6.10.: A) Homodyne interferometric detection employs a Michelson interferometer. The movable mirror is switched between two detection states (s sin φ and s cos φ)
from which near-field amplitude (s) and phase (φ) can be calculated. B) Heterodyne
interferometric detection scheme is based on an optical interferometer with an acoustooptic frequency shifter (AOM), a reflection path which couples to an AFM tip oscillating
at frequency f in z-direction, and a dual-output lock-in amplifier operating at the sum
frequency of either ∆ + nf [117].
71
(AOM) (Fig. 6.10 B). As for homodyne interferometric detection also in this case the
detected intensity consist of interference among three fields but with the difference that
the reference field is frequency shifted.
∗
Idetector = (EB + ER + EN )(EB∗ + ER∗ + EN
)
2
2
2
= |EB | + |ER | + |EN | + 2|EB ||EN | cos(φB − φN )
|
{z
} |
{z
}
(6.34)
+ 2|EB ||ER | cos(∆ωt + φR − φB ) +2|EN ||ER | cos(∆ωt + φR − φN )
|
{z
}
E ∗ = complex field conjugated to E, φ = phase, ∆ω = beat frequency
The intensity after the lock-in detection with a frequency ∆ω+nf is thus:
I = 2|EN ||ER | cos(∆ωt + φR − φN )
(6.35)
E ∗ = complex field conjugated to E, φ = phase, ∆ω = beat frequency
Using this kind of detection already at n = 2 pure near-field amplitude and phase contrast can be obtained. Experimentally the term 2|EB ||ER | cos(∆ωt + φR − φB ) is used
to adjust the interferometer alignment by observation of the beating frequency ∆ω with
an oscilloscope.
Recently a comparison between homodyne and heterodyne detection were published by
Gomez et al. [119].
6.3.4. Applications of SNOM
6.3.4.1. Aperture SNOM in the optical regime
In biological applications aperture-based SNOM is most frequently combined with fluorescence spectroscopy. A review on cell biological applications under ambient conditions
is given by de Lange et al. [121]. SNOM under physiological conditions is difficult
since common techniques for a probe-sample distance control are not as well suited for
operation in liquid as under ambient conditions. Among others individual fluorescent
molecules on the membrane of cells in solution were investigated [122], also the nuclear
envelope of Xenopus laevis oocytes were imaged in liquid [123].
SNOM has been applied to the characterization of organic thin films to correlate their
optical properties with sample morphology and to study phase separated polymer blends
[124, 125, 126, 127]. Information on nanoscale morphological texture of organic semiconductor thin films are for instance important for optimization of photovoltaic devices,
light-emitting diodes or field-effect transistors.
72
6.3.4. Applications of SNOM
6.3.4.2. Scattering SNOM in the optical regime
Raman spectroscopy, which is a complementary method to IR absorption spectroscopy
has also been combined with scattering-type SNOM. The group of Novotny demonstrated
raman spectra of free single-walled carbon nanotubes (SWNTs) on glass [128] and also
buried beneath a host dielectric medium [129]. But also with regard of biological research
different TERS experiments were reported. Among them spectral characterization of
bacterial surfaces [130, 131], detection of the membrane protein cytochrome c [132], and
investigation of single-stranded RNA [133].
6.3.4.3. Aperture SNIM
With regard to live cell imaging first aperture-based SNIMs were developed that can
carry out spectral and topographic imaging in liquid environment [134]. Those developments are very challenging due to mechanical problems with stability of the infrared
probes when vibrating in liquid. Experiments on different cell types were performed
using a free electron laser as light source [103, 107].
Also the potential of SNIM for studying the lateral organization of model membrane domains and clusters was demonstrated at a lateral resolution of about 100 nm by Generosi
et al. [135]. They reported on the spectroscopic investigation of DOPC multi-bilayers
deposited on glass in ambient conditions.
Chemical mapping of patterned polymer resist (poly(methylacrylate acid)(PMAA)) was
demonstrated by Dragnea et al. [136]. Characterization of mesoscale structures of
thin film polymers were performed in the C-H stretching region on polystyrene (PS)/
poly(ethyl acrylate)(PEA) [137]. In addition water vapor uptake in thin polymer films
was studied in a controlled environment chamber at a wavelength of 2.85 µm [138].
6.3.4.4. Scattering SNIM
Increasing interest in the ability of SNOM probing optical properties of materials with
nanometric resolution exists in material science.
High-resolution subsurface imaging provide a nondestructive, highly specific method for
mapping and chemical characterization of nanoscale objects within an embedding host
material. Subsurface imaging of gold islands 50 nm below a polymer surface was reported
by Taubner et al.[139] as well in the optical region (λ = 633 nm) as in the infrared regime
(λ = 10.7 µm) with a lateral resolution of <120 nm.
In addition studies on different implanted semiconductors were carried out by different
groups. Lahrech et al. investigated boron in silicon at a wavelength of λ = 10.6 µm
with a lateral resolution ∼ 400 nm [140]; Knoll et al reported on infrared imaging of
silicon samples with subsurface highly conducting stripes with a lateral resolution of 30
nm at λ approx. 10 µm [141]; Samson et al. were able to detect gallium implantations
in a silicon sample at λ = 3.22 µm [142]. Studying sub-surface local charge distributions
73
with nanometric resolution is of special interest with regard to miniaturization of components in microelectronics and characterization of conductivity in various solids such
as superconductors.
Furthermore the investigation of nanodomains of diblock-copolymer thin films has attracted much interest with respect to fundamental aspects and due to the technological
potential of block copolymers as functional materials for nanolithography, waveguides
and sensor applications. Akhremitchev et al. reported on investigation of polystyrenepoly(diemthylsiloxane) (PS-PDMS) diblock polymer [143]. Raschke et al. report on a
nanoscale surface analysis and identification of nanodomains on diblock copolymers at
a wavelength of 3.39 µm [144]. Taubner et al. investigated nanostructured PMMA/PS
polymer blend at wavelengths between 5.5 and 6 µm with a lateral resolution of 70 nm
[145].
With regard to biological applications only a few studies with s-SNIM were performed
so far but a further growth in this direction is expected.
Brehm et al. [146] were able to record an infrared near-field spectrum of a single tobacco
mosaic virus (TMV, 18 nm diameter, 400 nm length) on silicon in the amide region between 1600 to 1750 cm−1 with a lateral resolution of only 30 nm.
First experiments on supported lipid bilayers and vesicles with a sensitivity in the attomole regime were recently demonstrated proving the feasibility of s-SNIM on investigation of lipid membranes [147, 148]. Since s-SNIM does not require labeling of molecules
the investigation of phase transitions in lipid mixtures (lipid rafts) will be a very interesting topic for future studies.
Monolayer-sensitive chemical imaging was first demonstrated on microcontact printed
DNA stripes [149] and following on microcontact printed biotinylated alkylthiols [150].
6.3.4.5. s-SNOM in the THz regime
Moreover a combination of s-SNOM with THz radiation is a very promising concept.
Electromagnetic radiation at THz frequencies offers a variety of new possibilities for mapping and characterization of micro- or nanoelectronic devices, low-dimensional semiconductor nanostructures in material sciences or also cellular entities for biological research.
Cho et al. reported on first measurements with regard to high-frequency permittivity of
dielectric surfaces on submicrometer semiconductor structures [151]. Recently Huber et
al. investigated mobile carriers on single semiconductor nanodevices in a concentration
range of 10−17 to 10−18 cm−3 with a spatial resolution of about 40 nm at 2.54 THz
(λ = 118 µm) [152].
74
7. Preparation and characterization of
gold substrates
Contents
7.1. Materials and Methods . . . . . . . . . . . . . . . . . . . . . .
76
7.1.1. Sputter deposited gold surfaces . . . . . . . . . . . . . . . . .
76
7.1.2. Preparation of template stripped gold surfaces . . . . . . . .
76
7.1.3. SAM formation and cleaning with piranha solution . . . . . .
77
7.2. Results and Discussion . . . . . . . . . . . . . . . . . . . . . .
78
7.2.1. Comparison evaporated gold vs TSG . . . . . . . . . . . . . .
78
7.2.2. Cleaning with piranha solution . . . . . . . . . . . . . . . . .
80
7.3. Conclusion
. . . . . . . . . . . . . . . . . . . . . . . . . . . . .
81
75
Chapter 7. Preparation and characterization of gold substrates
Thin films of gold deposited on silica surfaces (e.g. glass, silicon wafer, mica) are common substrates for the formation of self-assembled monolayers (SAMs) from thiolated
molecules. Depending on the application and analysis method the choice of substrate
and the preparation process are influencing the morphology of the gold surface and
consequently also the quality of the resulting SAMs.
7.1.1. Sputter deposited gold surfaces
Gold coated silicon was purchased from Anfatec Instruments AG. A 125 nm thick layer
of gold was sputter deposited onto n(100) silicon (1.2 to 4.4 Ωcm) which had been primed
with an adhesion layer of 8 nm of titanium. Prior to use the substrates were cleaned
with hot piranha solution (3:1 mixture of 96% H2 SO4 and 30% H2 O2 ) for approximately
10 min, rinsed thoroughly with HPLC water and temporarily stored in HPLC water.
Piranha solution should be used freshly prepared. It is prepared by adding the peroxide
to the acid causing an exothermic reaction in which the solution can heat up to 120◦ C.
The cleaning efficiency decreases during cooling. The solution can be further heated
under thoroughly stirring to sustain its reactivity. Gold substrates should not be left
too long in the solution, since the roughness of the gold surface increases upon prolonged
exposure. Cleaning usually requires 5-15 min. A clean gold surface is hydrophilic [20, 21]
and completely wetted by water. The gold layer may even peel off, especially if no
adhesion layer between the gold and the substrate is present. When working with piranha
solution some security guidelines should be followed:
• work under a fume hood and wear protection equipment
• use glass ware and stainless steel tweezers
• piranha solution is extremely aggressive and can burn skin if not handled with care
• piranha solution can be explosive if the peroxide concentration is more than 50%
or the solution is mixed with organic solvents (e.g. acetone, isopropyl alcohol)
• hot piranha solution can violently boil splashing off the extremely acidic solution
After usage the solution should be left in an open container in order to cool down to
room temperature. Once cooled down the solution can be flushed down the drain with
copious amounts of water.
7.1.2. Preparation of template stripped gold surfaces
Ultraflat gold surfaces were prepared by a template-stripping method according to Stamou et al. [15]. Ordinary microscopy slides (BK7, Menzel GmbH) were cut in 5 x
15 mm2 pieces. Each piece of glass was first rubbed clean using a tissue soaked with acetone and was then exposed to hot piranha solution for 10-15 minutes. Subsequently they
were thoroughly rinsed with HPLC water and ethanol and dried in a stream of nitrogen.
The cleaned glass pieces were glued onto a p(100) silicon wafer (7-21 Ωcm) coated with
1500 Å gold (Anfatec Instruments AG) using the epoxy glue Epo-Tek 377 (Polytec PT
76
7.1.3. SAM formation and cleaning with piranha solution
GmbH). This two component glue consists of a resin and a hardener, which have to be
thoroughly mixed in equal parts by weight. After mixing a pipet tip was used to place
a small drop of glue on each glass slide. Then the glass slide was carefully placed on top
of the gold surface with the glue containing surface facing the gold. The amount of glue
is a very important issue. When using too less glue the glue is not wetting the whole
glass surface resulting in a poor adhesion of the gold to the glass. Surplus amounts of
glue will require large mechanical forces for the later removal of the glass pieces due to
a thick bulge of glue surrounding the outer part of the glass slide. After placing all glass
slides onto the gold surface the wafer is cured for 90 minutes at 150◦ C to cure the glue.
Finally the wafer is slowly cooled down.
Figure 7.1.: Preparation of template stripped gold. A support is glued onto a metal film
deposited on a flat template. The gold surface exposed to the glue has a roughness at the
nanometer scale whereas the stripped gold surface mimics the flatness of the template.
The resulting Si-Au-glue-glass ”sandwiches” (Fig. 7.1) could be stored as ”stripping
precursors” at least up to several months [14, 15] and be used whenever needed. Immediately prior to use the glass supported gold was mechanically stripped using a scalpel.
First, the blade of the scalpel was guided around one glass slide, and, afterwards it was
used to carefully lift-off the piece of glass at one edge. During the lift-off process caution
is advised because a shift of the glass slide on the silicon can easily cause scratches on
the gold surface.
7.1.3. SAM formation and cleaning with piranha solution
Cleaned sputter deposited gold substrates (see above) were incubated for 30 min in 1 mM
MCH in absolute EtOH and subsequently dried in a stream of nitrogen. After recording
IRRAS spectra the substrates were dipped for 1, 10 and 20 min in hot piranha solution,
thoroughly washed with water and ethanol and dried in a stream of nitrogen. Subsequently to further IRRAS measurements of the cleaned substrates they were incubated
in ethanolic solution of 1 mM ODT for 24 h and dried in a stream of nitrogen.
77
7.2.1. Comparison evaporated gold vs TSG
Surface morphology of sputter deposited plain gold surfaces and TSG surfaces were investigated using a modified Bermad 2000 AFM (Nanotec) in dynamic mode. The gain in
surface roughness when using TSG compared to a sputter deposited plain gold substrate
is illustrated in figure 7.2. Picture A and B display an area of 5 x 5 µm2 and C and
D are zooms into an area of 1 x 1 µm2 marked by black squares in the corresponding
image A and B.
For the sputter deposited gold a grainy surface structure was observed with grain dimensions of approximately 50-100 nm (Fig. 7.2, image C). Such a surface morphology
is characteristic for physical vapor deposited gold and reflects the nucleation-growth
mechanism of film formation when gold condenses from the vapor phase onto a substrate at room temperature. The height distribution ranges from 0 to 10.34 nm inside
Figure 7.2.: Dynamic mode AFM topographs of sputter deposited gold (left images
(A,C)) and TSG (right images (B,D)) surfaces. C and D are zooms (hardware) of the
upper topographs. The zoom area is indicated by the black squares.
78
7.2.1. Comparison evaporated gold vs TSG
an area of 25 µm2 (image A) and from 0 to 9.32 nm inside an area of 1 µm2 (image C).
Corresponding root-mean-square surface roughness (Rrms 1 ) are 1.2 nm over an area of
25 µm2 and 1.04 nm over an area of 1 µm2 area.
For preparation of TSG we followed the method introduced by Stamou et al. [15] who
used smooth silicon wafers as template instead of mica. Although these gold surfaces are
not as flat as mica (average roughness up to five times larger compared to mica [15]) the
advantage of this approach is derived from the mechanical properties of the silicon wafer,
which result in easier handling and ensure a complete separation between gold and template. Topographs B and D show a typical template stripped gold surface. Compared
to the sputter deposited gold TSG does not show gold grains but mimic the surface of
the silicon surface used as template. Black spots in image B are defects in the gold layer
likely caused by the stripping process and/or impurities on the silicon wafer (e.g. dust)
prior to metal deposition. The height of the TSG is ranging from 0 to 2.93 nm inside
an area of 25 µm2 (B - including the defects) and from 0 to 1.07 nm inside an area of
1 µm2 (D). The Rrms is 0.33 nm over an area of 25 µm2 and 0.16 nm over an area of 1 µm2 .
In comparison to sputter deposited gold TSG is approximately one order of magnitude
smoother. The criteria for selecting the type of substrate and method of preparation are
mostly dependent on the requirements of the technique for analysis and on the feasibility
of the preparation method. In SPM studies the surface roughness plays an important
role as already mentioned in the introduction. When the surface roughness is in the same
order of magnitude as the molecular structure of the adsorbed molecules both signals
intertwine and the dimensions of the adsorbed molecules get lost within the topographic
noise of the substrate. Therefore SPM experiments benefit from substrates as TSG
that offer very smooth surfaces. A shortcoming of the template stripping process is the
sample size. Small pieces are relatively easy to strip off but the larger the dimensions
the more difficult is the stripping process without damaging the gold surface. For other
applications such as IRRAS relatively large samples (several cm) are required to obtain
an adequate signal-to-noise ratio. In this case polycrystalline films are sufficient and
easier to prepare.
1
The surface flatness can be characterized by its root-mean-square roughness
Rrms
v
u
N
xN
u
X
= t[1/(N 2 − 1)]
(hmn − h̄2 )
(7.1)
mn=1
where NxN is the number of pixels, hmn is the height value of the pixel mn and h̄ is the mean height
of the pixel calculated from the NxN values and by its average roughness
RA =
N xN
1 X
| hmn − h | .
n mn=1
(7.2)
79
7.2.2. Cleaning with piranha solution
The cleaning efficiency of piranha treatment was tested on SAMs of MCH grown from an
ethanolic solution (Fig. 7.3, upper diagram). After SAM formation the substrates were
cleaned with hot piranha solution for 1, 10 and 20 min (Fig. 7.3, diagrams left side). In
order to demonstrated that SAM formation is not inhibited or diminished after piranha
treatment due to an increase of surface roughness or oxidization of the gold the cleaned
substrates were immersed into fresh solution of ODT for 24 h (Fig. 7.3, diagrams right
side).
All piranha cleaned substrates appear spectroscopically ”flat”, especially the previous
characteristic MCH peaks are lost. Remarkably even the 1 minute piranha cleaned substrate is free of any characteristic absorption features.
After 24 h of incubation in ODT solution each spectrum shows characteristic C-H stretching vibrations ranging from 3000 to 2800 cm−1 . Peak positions and band assignment are
given in table 7.1. The frequencies of the CH2 vibrational modes (νs at 2850 cm−1 and
Figure 7.3.: IRRAS spectra of a typical MCH SAM (upper diagram), MCH SAMs
after different incubation times in hot piranha solution (left side), and of subsequently
prepared ODT SAMs (right side).
80
7.3. Conclusion
peak center position
[cm−1 ]
2850
2877 and 2937
2917
2964
vibrational mode
CH2
CH3
CH2
CH3
sym. stretch
sym. stretch (FR)
antisym. stretch
antisym. stretch (ip)
Table 7.1.: Peak assignment for the IRRAS spectrum of ODT in the spectral region
from 3000 to 2800 wavenumbers. (FR = this band is split owing to Fermi resonance
interactions with the lower frequency antisymmetric CH3 deformation band; ip = in
plane)[29, 153].
νa at 2917 cm−1 ) confirm high density crystalline-like conformations for all three ODT
SAMs [29]. However, the intensity of the CH2 vibrational peaks of the sample that was
piranha cleaned for only 1 min are slightly reduced. This reduction in intensity might
be interpreted that way that not all of the gold surface is covered by ODT molecules
(submonolayer coverage). Since no shift of the CH2 peaks is observed the assembled
parts of the ODT film must be of crystalline order. The submonolayer coverage is most
likely due to spectroscopically invisible contaminants impairing full monolayer assembly. Therefore, based on the spectroscopic data a cleaning time of at least 10 min is
recommended.
7.3. Conclusion
TSG is five times smoother than sputter deposited gold. It is therefore favored as
substrate for scanning probe microscopy. However, polycrystalline gold is easier to
prepare in larger dimensions. It is therefore more suitable for IRRAS measurements
where the flatness does not play such an important role. When not freshly prepared
polycrystalline gold needs to be cleaned prior to use. Piranha solution is an effective
agent for cleaning gold surfaces. Even chemisorbed molecules such as thiols are removed
by piranha treatment.
81
8. Stamp fabrication and microcontact
printing (µCP) of ODT
Contents
8.1. Materials and Methods . . . . . . . . . . . . . . . . . . . . . .
84
8.1.1. Stamp fabrication . . . . . . . . . . . . . . . . . . . . . . . .
84
8.1.2. Stamp cleaning . . . . . . . . . . . . . . . . . . . . . . . . . .
85
8.1.3. Microcontact printing of ODT . . . . . . . . . . . . . . . . .
86
8.1.3.1. Preparation of ODT solution for stamp loading . . .
86
8.1.3.2. Microcontact printing of ODT . . . . . . . . . . . .
86
8.2. Results and Discussion . . . . . . . . . . . . . . . . . . . . . .
86
8.2.1. Stamp fabrication . . . . . . . . . . . . . . . . . . . . . . . .
86
8.2.2. Microcontact printed ODT patterns . . . . . . . . . . . . . .
87
8.3. Conclusion
. . . . . . . . . . . . . . . . . . . . . . . . . . . . .
89
83
Chapter 8. Stamp fabrication and microcontact printing (µCP) of ODT
A schematic overview of casting a PDMS stamp and µCP is presented in Figure 3.2
(page 24).
8.1.1. Stamp fabrication
Different commercially available AFM silicon calibration grids (MikroMasch) were used
as master. Figure 8.1 displays schemes of the calibration grids and Table 8.1 contains the
corresponding parameters. Before usage the desired calibration grid was cleaned with
hot piranha solution (7:3 mixture (v:v) of sulfuric acid and 30% hydrogen peroxide)
and thoroughly washed with HPLC water (for detailed handling of piranha solution
see chapter 7 (page 75)). For casting a stamp the master was fixed onto the bottom
of a plastic dish using a double-sided tape. A 10:1 ratio (w/w) mixture of SYLGARD
silicone elastomer 184 and SYLGARD silicone elastomer 184 curing agent (Dow Corning
Corporation) was thoroughly mixed for at least 15 minutes. Subsequently it was degassed
for approximately 30 min at room temperature using an exsiccator and a vacuum pump.
The prepolymer was carefully poured over the master until the petri dish was completely
filled. Afterwards it was cured for 2 days at room temperature to harden into a rubber-
Figure 8.1.: Scheme of AFM silicon calibration grids (Mikromasch) used for stamp
fabrication. The TGZ series have rectangular step patterns and the TGG calibration
grid has a triangular step pattern. Grid parameters are given in Table 8.1.
84
8.1.2. Stamp cleaning
name
lattice constant g
[µm]
TGZ 11
10
TGZ 04
3
TGG 01
3
height h angle α
[nm]
[◦ ]
1350 - 1650
900 - 1100
1800
70
Table 8.1.: Parameters of the silicon calibration grids (MikroMasch) used for stamp
fabrication. The TGG has triangular steps with curvature radii of less than 10 nm on
top. The TGZ series has rectangular steps. Schemes of both grid types are shown in
Figure 8.1.
like block. The elastomeric block was gently removed from the petri dish and peeled
away from the master. At that position where the master was placed a cavity was
formed in the PDMS block revealing an inverse replica of the features on the master:
raised regions of the master correspond to recessed regions in the PDMS. In order to be
used as a stamp the outer parts of the PDMS were cut away. Otherwise the patterned
PDMS area can not contact a substrate. The cuts have to be done with caution not to
harm the pattern. Figure 8.2 illustrates the cutting process.
Figure 8.2.: In order to create the stamp the calibration grid is glued into a petri dish
and the prepolymer is casted against it. After curing and removal of the PDMS from
the petri dish the excess PDMS has to be cut away (along the dashed line).
8.1.2. Stamp cleaning
Prior to use the stamp was cleaned in a 1:1 (v:v) mixture of water and absolute ethanol
in an ultrasonic bath. Therefore the stamp was put into an Eppendorf tube with the
stamping surface down. The Eppendorf tube was put with open cap into a 50 ml falcon
tube. Subsequently the falcon tube was filled with a 1:1 (v/v) mixture of HPLC water
and absolute ethanol until the small Eppendorf tube with the stamp inside was fully
covered. Then the closed falcon tube was sonificated for 15 minutes (if the stamp surface
was heavily contaminated up to 30 minutes). Finally the stamp was shortly rinsed with
ethanol and dried in a stream of nitrogen.
85
8.1.3. Microcontact printing of ODT
8.1.3.1. Preparation of ODT solution for stamp loading
Stamp loading was performed in a 5 mM ethanolic solution of ODT. The solution was
either freshly prepared or stored in a fridge at 4◦ C excluded from light. Because ODT
tends to precipitate (white flakes in the solution) it needs to be resolved before use.
Therefore the solution was heated in a water bath to 36◦ C for 15 minutes and was
shaken thoroughly. Alternatively it can be put 10 to 15 minutes into an ultrasonic bath.
8.1.3.2. Microcontact printing of ODT
The dry stamp was put, with the stamp surface down (to avoid contaminations of the
relief surface of the stamp), in the prepared Eppendorf tube filled with approx. 1 ml
of 5 mM ethanolic solution of ODT. When using the stamp for the first time it was
incubated over night. Otherwise it was immersed into ODT solution for 5 to 10 minutes.
In order to avoid precipitation of ODT during stamp loading the tube with the stamp was
put in a water bath at 36◦ C (very recommended for incubation over night). Subsequently
the loaded stamp was shortly rinsed with ethanol and predried in a stream of nitrogen for
at least 1 to 2 minutes. If the stamp was cleaned for longer than 10 minutes it was further
left in an Eppendorf tube for at least 1 to 2 hours (better over night) to dry completely
before usage. Otherwise deformations of the stamp pattern due to swelling or due to
residual solvent among the relief structures can occur. While preparing the substrate for
printing the stamp was temporary left in a dry and clean Eppendorf tube with the stamp
surface down. As substrates either gold coated n(100) silicon (Anfatec Instruments AG)
which was cleaned with hot piranha solution (7:3 mixture of 96% H2 SO4 and 30% H2 O2 )
for approximately 10 min or template stripped gold (for details see chapter 7, page 75)
was utilized. Printing was done by gently pressing the stamp to the gold surface by
hand. After a contact time of 90-120 s, the stamp was carefully peeled off from the
substrate and temporarily placed in an Eppendorf tube because further treatment has
to be given to the substrate as fast as possible to avoid contamination of the free gold
areas. Afterwards the stamp was thoroughly rinsed with ethanol to remove possible
contaminations resulting from the printing process and dried in a stream of nitrogen. If
further prints were done the stamp was again incubated for 1 to 2 minutes into ODT
solution and the process was repeated as described before. Finally the stamp was stored
in an Eppendorf tube, with the stamp surface down. The Eppendorf tube was well
closed and stored excluded from light at room temperature.
8.2.1. Stamp fabrication
Calibration grids as well as fabricated stamps were characterized with a reflected light microscope (WiTEC). Figure 8.3 B displays a light micrograph of calibration grid TGZ11.
86
8.2.2. Microcontact printed ODT patterns
Figure 8.3.: Scheme A depicts the dimensions of the calibration grid TGZ11 (MikroMasch). B shows a light micrograph of the calibration grid. Bright features correspond
to recessed regions. A light micrograph of the stamp fabricated from calibration grid
TGZ11 is presented in C. Bright features correspond to raised regions. D displays a
contact mode topograph of the stamp showing a step height of 1.5 µm and a lattice
constant of 10 µm.
Dark stripes correspond to raised regions with a width of 2.5 µm. The lattice constant
is 10 µm. Image 8.3 C depicts a PDMS stamp fabricated from this calibration grid. The
molded line pattern is homogeneous and defect free. Compared to image B the contrast
of image C is inverted meaning raised regions are brighter than recessed regions. A
contact mode topograph (BerMad 2000 AFM, Nanotec) of the stamp is shown in D.
The step height of the stamp relief pattern is 1.5 µm and the lattice constant is 10 µm.
8.2.2. Microcontact printed ODT patterns
In order to test the printing quality of the stamps ODT was printed on TSG surfaces.
Figure 8.4 depicts dynamic mode topographs (Bermad 2000 AFM, Nanotec) of those
patterns with corresponding line profiles. Recessed regions are displayed in dark.
Pattern B was printed with a stamp fabricated from a TGG01 calibration grid with
triangular steps. In accordance with the parameters of the calibration grid the lattice
constant of the printed lines is ∼3 µm. However, the printed ODT stripes are wider
than the width of calibration grid’s or stamp’s flat regions, respectively. Referring to
the dimensions of the calibration grid (angle of the triangle: 70◦ , height: 1.8 µm, lattice
87
Figure 8.4.: Dynamic mode topographs of printed ODT patterns (B, E) with corresponding height profiles (C, F). Printed ODT lines appear bright in the topographs.
Pattern B was printed with a stamp fabricated from a TGG01 grid (A). For printing
pattern E a stamp poured from a TGZ04 grid was used.
88
8.3. Conclusion
constant 3 µm) the width of the flat regions is approximately 0.5 µm:
width of flat regions = 3µm − 2 · (tan 35◦ · 1.8µm) = 0.48µm
(8.1)
Nevertheless the printed ODT lines are about 1 µm wide.
ODT lines in topograph 8.4 B were printed with a stamp casted against a TGZ04
calibration grid with rectangular steps. As expected the lattice constant of the printed
pattern is 3 µm. In this case, raised and recessed lines of the master pattern have a width
of 1.5 µm. Nevertheless the printed ODT stripes are again wider than the remaining
gold stripes where the stamp had no contact to the gold surface. The line profile shows
a width of approximately 2 µm for the ODT stripes.
Both printed patterns show a line broadening of approximately 0.5 µm. This is most
likely due to lateral diffusion of the ODT at the borders between surface contacting and
non-contacting regions of the stamp. Furthermore stamp deformations can occur caused
by the applied pressure during printing. Swelling of the stamp is unlikely because the
stamp was dried over night. Even if it was swollen due to cleaning in an ethanol/water
mixture and incubation in ethanolic ODT solution it should have gained back its initial
shape over night. In addition the long drying time prevents pattern distortion due to
residual solvent among the relief structures. Measured heights are in good agreement
with the theoretical total length of 2.5 nm for ODT.
8.3. Conclusion
Generally standard AFM calibration grids are well-suited and cheap masters for stamp
fabrication. A shortcoming is only the small outer dimensions of the calibration grids
(5x5 mm) which make handling of the resulting stamps more difficult. Another drawback
is the limited choice of shapes and patterns.
89
9. Experimental set-up of the
scattering scanning near-field
infrared microscope (s-SNIM)
Contents
9.1. Scattering scanning near-field microscope . . . . . . . . . . .
92
9.1.1. Set-up . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
92
9.1.2. AFM mode and lock-in detection . . . . . . . . . . . . . . . .
95
9.1.3. Beam alignment . . . . . . . . . . . . . . . . . . . . . . . . .
96
9.1.4. Precise sample positioning under the AFM tip . . . . . . . .
96
9.2. Carbon monoxide (CO) laser . . . . . . . . . . . . . . . . . .
97
9.2.1. Set-up and operation of the CO-laser . . . . . . . . . . . . . .
97
9.2.2. Electrode cleaning . . . . . . . . . . . . . . . . . . . . . . . .
100
91
Chapter 9. Experimental set-up of the scattering scanning near-field infrared microscope (s-SNIM)
9.1. Scattering scanning near-field microscope
9.1.1. Set-up
The basis of our scanning near-field infrared microscope is a BerMad 2000 AFM (Nanotec Electronics). In order to expand the AFM into a s-SNIM focussing of an additional
external radiation beam onto the AFM tip is required. Therefore it was necessary to
modify the conventional AFM head to gain additional space around the tip for illumination and detection of light. Details on the modification are given in reference [154].
Figure 9.1 presents the general set-up of the s-SNIM. The external beam is guided by a
mirror based beam path to the AFM. Since the beam path optics consist of mirrors it
allows using different radiation sources over a wide spectral range (provided that suitable detection optics are available). Currently we have integrated two different radiations
sources into the set-up: a tunable high-power tabletop opto-parametric oscillator (OPO)
which covers the spectral range from 3400 to 2600 wavenumbers (λ = 2.9-3.8 µm) and
a tunable liquid nitrogen cooled sealed-off CO-laser that emits infrared radiation from
2100 to 1600 wavenumbers (λ = 4.8-6.3 µm). Both beams are p-polarized (polarization perpendicular to the sample surface) because polarization plays a crucial role in
scattering scanning near-field microscopy. If the polarization of the incident light is not
parallel to the tip the local field between tip and sample surface is strongly attenuated
(see section 6.3.2.2 (page 64) and Fig. 6.8 (page 66)). Mirror M2 (Fig. 9.1), the lower
mirror of a beam lift, is mounted rotatable and enables a fast switching between both
radiation sources. Nevertheless, it is necessary to align the beam path when changing
the radiation source. After passing the lift the collimated infrared beam is expanded by
a factor of 4.57 using a telescope set-up consisting of two concave mirrors (d1 = 50.8 mm,
f1 = 100 mm, Fig. 9.1 M3; d2 = 76.2 mm, f2 = 457 mm, Fig. 9.1 M4). Since the aperture
of the mirror that focuses the beam onto the AFM tip is 25.4 mm the beam diameter is
delimited to 25 mm by a diaphragm (Fig. 9.1 D3) after the telescope. Then the beam
is focused by a 90◦ off-axis parabolic mirror (f = 101.6 mm, Fig. 9.1 M7) onto the cantilever with an ∼80◦ angle of incidence. Ideally the focal spot size is diffraction limited
and can be calculated using Equation 9.1:
2w = 1.22
f ·λ
d
(9.1)
w = beam radius, f = focal length, λ = wavelength, d = diameter of focussing element
Accordingly, the diameter of the focal spot ranges from ∼14.5 µm at λ = 3 µm to
∼30 µm at λ = 6 µm.
As SNIM probes commercial gold-coated cantilevers (NSC16/Cr-Au, MikroMasch) with
a tip curvature radius of <50 nm and a typical resonance frequency of 170 kHz were
employed. The gold-coating provides a good polarizability (Au = -1240 + ι3484 at
1711 cm−1 [155]) and therefore a strong local field enhancement.
Finally, the scattered radiation from the AFM tip in local interaction with the sample surface is collected by a calcium fluoride lens (CaF2 , d = 25.4 mm, f = 40 mm)
92
9.1.1. Set-up
Figure 9.1.: Set-up of the s-SNIM. M1 = kinematic mirror, D = diaphragm,
M3 and M4 = concave mirrors, M5 and M6 = plane mirrors, M7 = 90◦ offaxis parabolic mirror, MCT = Mercury Cadmium Telluride detector
93
and focused onto a liquid nitrogen cooled Mercury Cadmium Telluride (MCT) detector
(J15D12-204-S01M-60, Judson Technologies). The experimental set-up uses a 90◦ scattering scheme detection (i.e. the detected light is collected from a solid angle centered
90◦ off the incident radiation). Lens and MCT detector are positioned on separated
x-y-z translation stages in order to facilitate the fine adjustment. Figure 9.2 displays
the AFM and the detection unit enlarged.
Figure 9.2.: Set-up of the AFM and s-SNIM detection unit
94
9.1.2. AFM mode and lock-in detection
9.1.2. AFM mode and lock-in detection
SNIM experiments are performed in dynamic mode, which means that the tip oscillates vertically (perpendicular to the sample surface) with its resonance frequency f.
Due to the height oscillation of the cantilever the scattered radiation from the tip is
amplitude-modulated. However, the total signal intensity received by the detector contains additional to this AC component a DC component, i.e. non-modulated background
radiation (e.g. scattered light from the sample surface). The detected signal is preamplified (PA-101, Judson Technologies) and subsequently phase-sensitive filtered by lock-in
amplification which allows effective suppression of the non-modulated stray light. The
lock-in amplifier (SR 844, Stanford Research) is able to work in the 25 kHz to 200 MHz
range and is thus suitable to utilize the resonance frequency (f) of many AFM cantilevers
and their first harmonics (2f) as reference frequency. Unfortunately this lock-in amplifier
does not allow to record higher harmonics than twice the resonance frequency. Therefore detection at twice the resonance frequency of the cantilever oscillation (2f ∼340
kHz) is performed. Detection at higher harmonics yields an additional suppression of
modulated background signal, meaning the modulated signal that is not scattered from
the tip apex but from the rest of the cantilever without containing any near-field signal.
With increasing harmonic order the enhancement of the near-field signal over the background rises and the signal-to-background ratio is improved (for detailed description see
section 6.3.3 (page 67)).
√ After amplification and noise reduction the phase independent
amplitude signal R = X 2 + Y 2 is recorded using the lock-in amplifier. Recording only
the X output channel or Y output channel can induce phase artifacts in the near-field
image. Alterations in the mechanical phase of the cantilever due to changing surface
properties influence the phase of the optical modulation. If the signal channel phase
changes (but not its amplitude) the Y output as well as the X output will change. One
will decrease and the other one will increase. Consequently the vector magnitude R remains constant. When using lock-in detection the relation between scan parameters (i.e.
scan frequency and number of scan points) and time constant of the lock-in amplifier
has to be considered. For selecting an adequate time constant one has to know the time
per pixel which can be calculated from Equation 9.2.
time per pixel in ms =
1000
number of pixels per line · 2 · scan frequency (Hz)
(9.2)
If the time constant is larger than the time per pixel the image becomes blurred because
the output signal is averaged over more than one pixel. However
√ a time constant (τ )
that is too short leads to a poor signal-to-noise ratio (SNR ∼ τ ). Changes in the
input signal take a couple of time constants to be reflected at the output. This is mainly
because the RC low pass filter(s) at the input of the lock-in amplifier require(s) a certain
amount of time to settle to its final value (I(t) = Imax e−t/τ = Imax e−t/RC ). Therefore the
time constant reflects how slowly the lock-in output responds to changes at the input.
The delayed response is equivalent to an output smoothing. In praxis a time constant
that is less than one third of the time per pixel is recommended.
95
9.1.3. Beam alignment
The beam alignment is very crucial. In order to facilitate the alignment of the invisible
infrared light along the beam path a Helium-Neon laser copropagates with the infrared
beam and visualizes the path. The Helium-Neon beam is coupled into the beam path
using a kinematic mirror (Fig. 9.1 M1). Both beams are guided through two diaphragms
with a distance of 2 m to superimpose their beam path onto one beam axis (Fig. 9.1 D1
and D2). During alignment it is important to avoid optical aberrations such as astigmatism. Aberrations result in a large deformed focus onto the cantilever that is no more
diffraction limited. Consequently a larger area of the cantilever is illuminated leading to
a higher background signal. In general it is advisable to align the beam in a way that
it always hits the center of the optics. Especially in case of the focussing mirrors, i.e.
telescope mirrors and 90◦ off-axis mirror, this is crucial to avoid a beam deformation.
Furthermore the beam axis should be ideally perpendicular to the optical axis of the
components. Concerning the telescope (Fig. 9.1 M3 and M4) this is not realizable but
the angle of incidence should be kept as small as possible. In order to align the beam axis
on the 90◦ off-axis mirror (Fig. 9.1 M7), which focuses the beam onto the cantilever, mirror M5 and M6 are used. In addition, to keep the focus as small as possible an accurate
focussing onto the AFM tip is required. To this end the mirrors M6 and M7 (Fig. 9.1)
are mechanically combined on a special x-y translation stage allowing to move both
mirrors at the same time and change the distance between off-axis mirror and cantilever
(Fig. 9.1 M6 and M7 x-translation stage). On the other hand the off-axis mirror can be
moved independently from mirror M6 moving the beam along the cantilever arm (from
back to forth (Fig. 9.1 M7 y-translation stage)). Furthermore the off-axis mirror is fixed
in a rotational mount (360◦ ) providing an additional degree of freedom for aligning the
beam onto the AFM tip. Focussing onto the tip can be controlled by carefully observing
the silhouette of the AFM tip. The easiest way is to search the silhouette when the
tip is out of focus and to approach the focus slowly. With decreasing distance between
focus and tip the silhouette becomes larger and in close proximity to the focus a mirror
image of a second tip appears opposite to the original one. Silhouette and mirror image
approach each other until they finally ”melt” together and many diffraction patterns
appear. At this point the tip is in the focus.
In the end, the collecting CaF2 lens and the detector needs to be aligned. In the dark,
one can see a very weak scattering spot of the Helium-Neon laser that is used to prealign
lens and detector. The lens is fixed in that way that the weak spot hits the middle of the
lens and the outcoming beam has a small round focal size and shows no deformations.
Then the detector is aligned to that point. Afterwards the fine alignment is performed
using the infrared radiation after approaching the AFM tip to a sample.
9.1.4. Precise sample positioning under the AFM tip
In order to align a sample to a defined location (e.g. a scratch mark) under the AFM
tip a stereo-microscope (MZ6, Leica) with a flexible light guide is fixed over the AFM
head. Since the cantilever as well as the sample surface have to be within the focal
96
9.2. Carbon monoxide (CO) laser
plane of the stereo-microscope the best way is to approach first the sample until it is in
contact with the tip and then withdraw it slightly. The glass prism which guides the red
diode laser beam onto the cantilever is removed to have a free view onto the cantilever
and the stereo microscope is focused onto the sample plane. Because the AFM sample
holder does not have a mechanical translation stage the positioning of the sample is
done carefully by hand. One can either move the sample itself or the whole piezo mount
which rests on the stepper motors. Moving the piezo mount is easier but the range is
limited to some millimeters. Anyway the positioning is very critical due to the close
proximity between sample surface and tip and has to be done with great care to avoid
damaging the sample or the tip. Finally the prism is reinstalled and a realignment of
the red diode laser beam onto the AFM tip is required.
9.2. Carbon monoxide (CO) laser
All s-SNIM experiments in this thesis were carried out with a sealed off CO-laser. Consequently only this infrared radiation source is described in detail. A detailed description
of the OPO is found in [156, 142].
The CO-laser is a molecular gas discharge laser and one of the major sources in the
mid infrared due to its potential high power and broadband emission. It was originally developed in 1965 by C.K.N. Patel [157] and is based on laser transitions between vibrational-rotational transitions in the electric ground state of the CO molecule.
Pumping into higher vibrational states occurs via electron excitation and anharmonic
vibration-vibration energy transfer (anharmonic VV-pumping) [158]. Initially the laser
provided several hundred laser lines in the wavelength region between 4.8 and 8.3 microns. In 1971 Freed [159] reported on the first sealed-off CO-laser allowing the use of
expensive CO isotopes which enable an increase in the number of available laser lines.
About 20 years later Urban et al. [160, 161] extended the emission range to 2.6-4 µm
by realizing laser emission for vibrational overtone transitions (∆v=2) in CO.
A review of CO-laser development and its spectroscopic application has been given by
Urban [162].
9.2.1. Set-up and operation of the CO-laser
Figure 9.3 presents a block diagram of the home-built sealed off nitrogen cooled CO-laser
used for s-SNIM experiments. The laser consists of a glass tube (i.d. 18 mm) immersed
in a sand vacuum which is surrounded by a liquid nitrogen dewar and a vacuum chamber. Nitrogen cooling is required to achieve low plasma temperatures resulting in a high
density of laser lines and high output power. At cooling temperatures below 110 K
(-163◦ C) laser gas components or chemical reaction products can freeze out. Therefore
laser glass tube and liquid nitrogen reservoir are separated by a sand-helium mixture
(Fig. 9.3, sand vacuum). The sand-helium mixture allows adjusting the thermal conductivity by variation of the residual helium pressure (typically ∼ 5 · 10−2 mbar). An
additional heating wire, in combination with a temperature sensor on the outside of the
97
Figure 9.3.: Set-up of a liquid nitrogen cooled sealed off CO-laser.
discharge tube, is used for controlling the cooling temperature. Optimal laser performance is achieved at temperatures around 133 K (-140◦ C).
At both ends the laser glass tube is sealed with two CaF2 windows mounted at Brewster’s
angle. Thereby the windows can serve as polarizing filter. High vacuum glue TorrSeal
from Varian GmbH is used to mount the CaF2 windows and the electrode branches. If
necessary this glue can be removed by carefully heating with a heat gun.
Next to the cathode a gas intake valve connects the laser tube with a gas mixer and
a turbo pump which allows archiving a base pressure of 2x10−5 mbar. Before laser
operation the gas mixer is filled with laser gas up to a pressure of 19 to 20 mbar (corresponding to 1.9 to 2 V at the baratron). Subsequently the valve between gas mixer and
laser tube is opened and the gas expands inside the laser tube. Then the valve is closed
again and the gas mixer is evacuated. In Table 9.1 the composition of our standard laser
gas mixture, which is purchased from Messer Griesheim GmbH (Krefeld, Germany), is
listed. Laser plasma excitation occurs due to a DC gas discharge. A current regulated
power supply provides high voltage to a hemispherical anode out of VA-steel associated
with a cylindrical nickel cathode at ground potential. Typical discharge conditions are
component
debit
credit
[Vol-%]
[Vol-%]
Xenon 4.0
3
2.92
Carbon monoxide 1.8
6
5.89
Nitrogen 5.0
10
9.02
Helium 6.0
remainder remainder
Table 9.1.: Standard CO-laser gas composition (production date: 04/2003)
98
9.2.1. Set-up and operation of the CO-laser
I = 10-20 mA and U = 15-16 kV.
The population inversion required for laser operation is achieved in three steps. First,
accelerated free electrons collide with CO molecules and excite them into low vibrational
states (v≤8):
CO + e− → CO− → CO∗ + e−
(9.3)
Pumping into higher vibrational states (up to v = 40) occurs via anharmonic vibrationvibration energy transfer (VV-pumping). Thereby excited CO molecules collide with
each other either in an exothermic or endothermic way.
CO(ν) + CO(ω > ν) → CO(ν − 1) + CO(ω + 1) + ∆E
(9.4)
CO(ν) + CO(ω > ν) → CO(ν + 1) + CO(ω − 1) − ∆E
(9.5)
or
Beside this VV-pumping a deactivating vibrational-translational relaxation (VT-relaxation)
can occur. The collision partner is any molecule within the laser gas (i.e. CO, He, Xe
or N2 ).
CO(ν) + M → CO(ν − 1) + M
(9.6)
At low temperatures the exothermic process 9.4 dominates over the deactivating endothermic reaction and the VT-relaxation. Therefore at plasma temperatures between
120-180 K a quasi equal population distribution for the vibrational states v=8-35 occurs.
The required population inversion is achieved due to the splitting of vibrational states
into rotational states. Contrary to the VT-relaxation the rotational-translational relaxation is not inhibited at low temperatures due to their lower energies. For this reason a
temperature dependent Boltzmann distribution of the rotational states occurs. Taking
into account also the degeneration of the rotational states results in a partial population
inversion among single rotational-vibrational states. The partial population inversion
for p-transitions (∆J = -1) of the CO molecules within the region of equally populated
states leads consequently to a laser activity.
The laser resonator has a total length of 1.5 m and is flushed with nitrogen during
operation to avoid water absorption. It consists of a gold coated concave mirror with
5 m radius of curvature and a gold coated reflection grating in Littrow configuration
(250 lines/mm). Out coupling of the zeroth order reflection of the grating occurs via a
retroreflector. Tunability is achieved by rotating the grating with a micrometer screw
either by hand or by a stepper motor. A shortcoming of the Littrow configuration is
that rotation of the grating results in displacement of the out coupled beam. Therefore
the beam alignment on the two diaphragms within the optical path of the SNIM set-up
(Fig. 9.1 D1 and D2) has to be checked after every change in grating position.
The laser is line tunable in the region from 2100 to 1600 wavenumbers (λ = 4.8-6.3 µm)
on discrete lines with an output power of typically ≥1 W. CO transitions and laser
lines are listed in Table C (page 185). For the s-SNIM experiments the output power
is attenuated to approximately 250 mW. The laser provides more than 100 lines with a
typical frequency separation of 3 cm−1 between adjacent laser lines. Due to the sealed
off construction the use of additional high cost CO isotopes (12 C16 O, 13 C16 O, 12 C18 O) is
99
possible, increasing the number of available laser lines to about 400. This corresponds
to an average spacing of 1.2 cm−1 between laser lines, which provides a good coverage
of the amide region. The linewidth is about 150 kHz. Figure 9.4 shows three emission
spectra of the CO-laser. A typical emission spectrum recorded with the standard conditions used for our experiments within the amide region is presented in diagram A. In
order to investigate organometallic infrared dyes with s-SNIM (chapter 12 (page 159))
a shift to higher frequencies (lower vibrational transitions) is necessary. The spectral
emission range of the CO-laser is influenced by two factors: (i) the plasma temperature
(i.e. discharge conditions) and (ii) the gas composition. In general low current and low
CO-content favors low plasma temperatures and low vibrational transitions. High vibrational bands require a high CO content and high currents (for a detailed explanation
see reference [163]). When operating the laser at a low current of 9 mA only a few
additional vibration bands occur in the high frequency region (spectrum B). Whereas
a decrease in CO content (diluting the laser gas with Helium while retaining the total
pressure constant) results in emission bands around 1950 wavenumbers (spectrum C).
9.2.2. Electrode cleaning
Carbonic deposits at the electrodes which appear especially at high CO concentrations
give rise to discharge instabilities. For cleaning the laser is filled with a 3:1 mixture of
helium and oxygen (He pressure: 3-3.6 mbar (corresponding to 0.3-0.36 V at the baratron and O2 pressure: 1-1.2 mbar (corresponding to 0.1-0.12 V at the baratron)). Then
a high voltage is applied for 5-10 min.
For removal of tenacious impurities the laser has to be opened at either the anode or the
cathode. Opening and closing of the laser always bears the risk of introducing a leakage
at the glued areas at the laser tube due to mechanical stress. Therefore special caution
is advised when handling the electrode branches.
Anode cleaning is performed in citric acid for 10-15 min and can be improved by sonification. Subsequently the anode is washed thoroughly with water and finally wiped with
a soaked tissue of ethanol or isopropanol. If the anode surface is blunted it needs to be
polished in the work shop. A proper connection between the high voltage supply and
the anode is important. Otherwise when applying a voltage a loud ”cheep” resounds
and the discharge jitters.
Cleaning the cathode is more difficult because it cannot be removed from the laser. To
this end the laser is opened at the (gas intake) and the cathode is cleaned carefully with
cotton swabs and a soft brush soaked with ethanol and/or isopropanol.
100
9.2.2. Electrode cleaning
Figure 9.4.: CO-laser spectra recorded at different discharge conditions. Missing lines
within the spectra are mostly due to water absorption in the resonator. By changing the
discharge conditions the spectrum can be shifted to lower or higher vibrational bands.
101
10. Characterization of
microstructured monolayers of
biotinylated alkylthiolates by
s-SNIM
Contents
10.1. Introduction
. . . . . . . . . . . . . . . . . . . . . . . . . . . . 104
10.2. Materials and Methods . . . . . . . . . . . . . . . . . . . . . . 104
10.2.1. Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
104
10.2.2. Preparation of homogeneous SAMs for FTIR measurements .
104
10.2.3. Preparation of microstructured SAMs . . . . . . . . . . . . .
105
10.2.4. Sample preparation for volume FTIR spectra . . . . . . . . .
105
10.2.5. Scanning electron microscopy (SEM) . . . . . . . . . . . . . .
105
10.2.6. Fourier Transform Infrared (FTIR) Spectroscopy . . . . . . .
105
10.2.6.1. Infrared Transmission Absorption Spectroscopy
. .
105
10.2.6.2. Infrared Reflection Absorption Spectroscopy (IRRAS) 106
10.2.7. Scattering scanning near-field infrared microscopy (s-SNIM) .
106
10.3. Results and Discussion . . . . . . . . . . . . . . . . . . . . . . 106
10.3.1. Scanning Electron Microscopy (SEM) . . . . . . . . . . . . .
106
10.3.2. Comparison of a volume and IRRAS spectrum of BAT . . . .
107
10.3.3. IRRAS spectra of BAT and ODT SAMs . . . . . . . . . . . .
108
10.3.4. s-SNIM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
111
10.3.4.1. Theoretical predictions . . . . . . . . . . . . . . . .
115
103
Chapter 10. Characterization of microstructured monolayers of biotinylated alkylthiolates by s-SNIM
10.1. Introduction
The results presented in this chapter are published as ”Chemical Imaging of Microstructured Self-Assembled Monolayers with Nanometer Resolution” [150]. Experiments on
microstructured monolayers of 1-octadecanethiolate (ODT) and biotinylated alkylthiolate (BAT) demonstrate the ability of s-SNIM to provide infrared spectroscopical information of thin organic films at the nanoscale. At a wavelength of 5.85 µm a lateral resolution of ∼90 nm was achieved comparable with a diffraction limited resolution of λ/60.
The detection limit was 5 x 10−20 mol corresponding to 27 attogram or 30.000 molecules
of BAT.
10.2.1. Materials
Biotinylated alkylthiol (BAT) was synthesized in the group of Prof. Dr. A. Terfort
(University of Frankfurt). 1-octadecanethiol (ODT, Aldrich, 98%) was purchased from
Sigma. Molecular structures of both molecules are presented in Figure 10.1. Ethanol
was used in p.a. quality.
Figure 10.1.: Bond-line structures of BAT and ODT
10.2.2. Preparation of homogeneous SAMs for FTIR measurements
Sputter deposited gold on silicon (Anfatec Instruments AG) was used as substrate (see
chapter 7 (page 75)). Prior to use substrates were cleaned with hot piranha solution (7:3
mixture of 96% H2 SO4 and 30% H2 O2 ) for approximately 15 min (detailed description
see section 7.1.1 (page 76)). Homogeneous SAMs were prepared by immersing clean gold
substrates into 1 mM ethanolic solutions of the pure thiols for approximately 120 min
104
10.2.3. Preparation of microstructured SAMs
at room temperature. Afterwards the substrates were rinsed with absolute ethanol and
dried in a stream of nitrogen.
Micropatterns of ODT and BAT were prepared by using microcontact printing. PDMS
stamps used in these experiments were fabricated in the group of Prof. Dr. A. Terfort
(University of Frankfurt). Electron microscopic images of a stamp evaporated with a
thin graphite layer are presented in Figure 10.2. Microcontact printing was performed
as described in chapter 8 (page 83). Briefly, prior to use, the stamp was cleaned and
subsequently loaded with ODT by immersion in a 5 mM ethanolic solution of ODT.
Printing was done by gently pressing the dried stamp on a clean gold substrate. After
a contact time of 120 s the stamp was carefully peeled off. Immediately after printing
the substrate was immersed into a 1 mM ethanolic solution of BAT in order to fill the
remaining bare gold areas with BAT. After 80 min. the substrate was thoroughly rinsed
with absolute ethanol and dried in a stream of nitrogen.
10.2.4. Sample preparation for volume FTIR spectra
For preparation of a BAT-KBr pellet, 200 mg of dry KBr were carefully pulverized and
thoroughly mixed with 2 mg of BAT. Afterwards the powder was filled into a hydraulic
press to form a pellet.
10.2.5. Scanning electron microscopy (SEM)
Scanning electron microscopy was carried out at the central SEM of the Ruhr-University
Bochum by Dr. R. D. Neuser. Prior to imaging the PDMS stamp was coated with a
thin layer of carbon to make it electrically conductive.
10.2.6. Fourier Transform Infrared (FTIR) Spectroscopy
FTIR measurements were carried out by Dr. Ch. Grunwald on a commercial Bio-Rad
FTS-3000 Excalibur spectrometer. Data are presented with his permission.
10.2.6.1. Infrared Transmission Absorption Spectroscopy
FTIR spectra on KBr pellets were recorded in transmission configuration using a DTGS
detector. After mounting the reference pellet (sample pellet) the sample chamber was
purged with dry air for 5 min. Spectral resolution was set to 4 cm−1 and 128 scans
were accumulated, averaged, and transformed by using a triangular apodization. For
reference a spectrum of a pure KBr pellet without thiol was measured.
105
10.2.6.2. Infrared Reflection Absorption Spectroscopy (IRRAS)
IRRAS spectra were recorded in grazing incident single-pass configuration with an incident angle of ∼80◦ against the normal of the surface. After inserting the reference or
sample into the sample chamber the spectrometer was purged with dry air for 5 min.
Spectra were recorded at room temperature with a resolution of 4 cm−1 . For the reference measurements 2048 scans were summed up and the sample measurements were
carried out with a coaddition of approximately 2000 scans. It was stopped when the
water bands disappeared. Interferograms were fourier transformed by using a triangular
apodization.
For reference a perdeuterated docosanthiol SAM was used. Referencing to a perdeuterated SAM instead of a bare gold substrate offers a clean, well defined and reproducible
reference surface. Perdeuterated alkanethiolates are well-suited as reference since they
prevent adsorption of contaminants onto the surface due to their hydrophobic properties
and the deuteration avoids spectroscopic overlap with the C-H vibrations.
10.2.7. Scattering scanning near-field infrared microscopy (s-SNIM)
s-SNIM measurements were carried out as described in chapter 9 (page 91) under ambient
conditions. Images were recorded with a scan rate of 0.5 Hz per line and a time constant
of 3 ms at the lock-in amplifier.
10.3.1. Scanning Electron Microscopy (SEM)
Scanning electron microscopy was applied to a PDMS stamp coated with a thin layer of
graphite, and a sample with a patterned SAM of ODT and BAT.
Figure 10.2.: SEM photomicrographs of a PDMS stamp coated with a thin layer of
carbon. The stamp relief pattern consists of ∼1.9 µm wide raised lines separated by
∼1.3 µm (top view A) with a height of ∼1 µm (side view B).
106
10.3.2. Comparison of a volume and IRRAS spectrum of BAT
Figure 10.3.: SEM photomicrographs of the structured substrate with 2.5 µm wide ODT
lines (bright lines) separated by 1.5 µm broad BAT stripes (dark lines).
Figure 10.2 A presents a SEM top view of a coated PDMS stamp with a line pattern.
The relief pattern consist of raised lines with a width of ∼1.9 µm separated by ∼1.3 µm
wide recessed regions. A slightly magnified side view of the stamp relief pattern (figure 10.2 B) shows a height of ∼1 µm for the raised stripes.
In Figure 10.3 two electron micrographs of a laterally structured SAM of ODT and BAT
on a sputter deposited gold substrate are displayed. ODT was printed using the stamp
shown in Figure 10.2. Refilling the remaining gold areas with BAT results in a periodic
pattern of ∼2.5 µm wide lines of ODT separated by ∼1.5 µm broad lines of BAT.
Comparing the width of the printed ODT lines and line width of the stamp a broadening
of the printed lines of about 0.5 to 1 µm is observed in SEM as well as in AFM topography
(Fig. 10.6). As already discussed in chapter 8 (page 83) this broadening can be caused
by different effects such as lateral diffusion of the molecular ink or undefined loading
forces during printing.
10.3.2. Comparison of a volume and IRRAS spectrum of BAT
Figure 10.4 displays an IRRAS spectrum of a BAT SAM in comparison with a FTIR
volume spectrum of a KBr pellet containing BAT. For comparison the surface spectrum
is multiplied with a factor 103 because the signals of thin films in IRRAS configuration
are much weaker due to the small amount of molecules. In diagram A an overview over
the whole spectral range from 3600 to 950 cm−1 is displayed. Diagram B shows the
regions from 3600-2700 cm−1 , corresponding to the O-H and C-H stretching vibration
region, and the lower frequency region from 1900-950 cm−1 , containing the amide bands,
enlarged. A detailed assignment of the vibrational modes is done in Table 10.1. The
peak at 1640 cm−1 in the volume spectrum that does not appear in the surface spectrum
is not a molecule specific peak that is missed due to the surface selection rule. This peak
arises from water inclusions in the KBr pellet due to the hygroscopy of alkalihalogenides
[153]. The very broad mode at 3300 wavenumbers in the BAT spectrum might be assigned to an amide A vibration (N-H stretching vibration in resonance with amide II
107
overtone vibration) [164]. An alternative explanation would be intermolecular hydrogen
bonds [164]. The ”noise” in the ODT spectrum in the region between 1800 cm−1 to
1600 cm−1 is caused by atmospherical water vapor [153].
In SAMs the molecules can be crystalline-like ordered while in a volume sample (mixed
KBr pellet) a random distribution exists. Therefore a volume sample allows the excitation of all possible vibrational modes of a molecule whereas for a surface spectrum the
surface selection rule has to be taken into account. For this reason volume and surface
spectra of the same substance can differ from each other, e.g. in shape of vibrational
modes (disorder result in peak broadening) or missing peaks in the surface spectrum.
In case of BAT the volume and surface spectra are very similar and no missing vibrational modes are observed in the surface spectrum. From this can be concluded that the
BAT molecules are assembled in a disordered monolayer resulting in different molecular
orientations.
10.3.3. IRRAS spectra of BAT and ODT SAMs
Figure 10.5 presents IRRAS spectra of monolayers of ODT and BAT. In diagram A an
overview over the whole spectral range from 3600 to 950 cm−1 is shown. Diagram B
displays the interesting regions from 3600-2700 cm−1 , corresponding to the O-H and C-H
stretching vibration region, and the low frequency region, containing the characteristic
amide region from 1900-950 cm−1 enlarged. Peak assignment is given in Table 10.1.
With respect to biomolecules, especially proteins and polypeptides, the spectral range
from 1500 to 1800 wavenumbers is well known as the amide vibration region. BAT
shows two prominent peaks in this region. One at 1550 cm−1 which is assigned to the
amide II vibrations consisting of ∼60 % N-H in-plane bending vibrations and ∼40 %
C-N stretching vibrations. The other prominent mode is centered at 1711 cm−1 and
caused by the C=O stretching vibration of the ureido group (R-HN-(C=O)-NH-R). The
weaker shoulder at 1650 cm−1 is an amide I vibration assigned to the two amide groups
in the chain of the BAT molecule. The negative peaks between 2000 and 2200 wavenumbers correspond to symmetric and antisymmetric CD2 and CD3 stretching vibrations of
the perdeuterated docosanthiol-SAM that was used as reference. The negative bands
in the region from 2300 to 2390 cm−1 correspond to the P- and Q-branch of the CO2
inside the spectrometer chamber. The long frequency range of the spectra contains the
C-H stretching vibrations for both molecules which appear sharper for the ODT in comparison to BAT. This is due to the better ordering of the ODT and a more liquid like
ordering of the BAT which is in good agreement with the conclusion from the comparison
of the surface and volume spectrum of BAT. Another indication for the disordering of
the BAT are the frequencies of the CH2 modes which are shifted to higher wavenumbers
compared to the corresponding peak locations of the ODT SAM (see Table 10.1).
108
10.3.3. IRRAS spectra of BAT and ODT SAMs
Figure 10.4.: Normalized IRRAS spectrum of BAT (black line) in comparison to an
FTIR volume spectrum of BAT (red line). Diagram A shows the frequency range from
3600 to 950 cm−1 . Diagram B displays the low frequency region (1900 to 950 cm−1 )
containing the characteristic amide region (1700 to 1500 cm−1 ) and the region of the
C-H and O-H stretching vibration (3600 to 2700 cm−1 ) enlarged. Peak assignment is
given in Table 10.1
109
Figure 10.5.: IRRAS spectra of BAT (black line) and ODT (blue line) monolayers on
gold. Diagram A shows the frequency range from 3600 to 950 cm−1 . Diagram B displays
a zoom in the low frequency region (1900 to 950 cm−1 ) with the characteristic amide
bands (1650 to 1550 cm−1 ) and the region of the O-H and C-H stretching vibration
(3600 to 2700 cm−1 ). Peak assignment is given in Table 10.1.
110
10.3.4. s-SNIM
[cm−1 ]
2965
2919, 2927
2878, 2936
2850, 2860
2300 - 2390
2220
2195
2089
1711
1650
1550
1464
1381
1267
1141
vibrational mode
CH3 antisym. stretch. (ip)
CH2 antisym. stretch.
CH3 antisym. stretch. (FR)
CH2 sym. stretch.
CO2
CD3 antisym. stretch.
CD2 antisym. stretch.
CD2 sym. stretch.
N−(C=O)−N
amide I
amide II
N−(C=O)−N antisym. stretch.
CH2 wags
CH2 wags and twists
C−N stretch. + N−(C=O)−N antisym stretch.
CH2 wags and twists
C−O−C antisym. stretch.
Table 10.1.: Peak assignment for the IRRAS spectra of BAT and ODT in Figure 10.5 in
the spectral region from 3000 to 1000 wavenumbers. A full assignment for BAT based on
theoretical calculations can be found in [165]. (FR = this band is split owing to Fermi
resonance interactions with the lower frequency antisymmetric CH3 deformation band;
ip = in plane)
10.3.4. s-SNIM
Figure 10.6 presents simultaneously recorded topography (A) and near-field (B) images
of a micropatterned SAM of ODT and BAT at an IR frequency of 1711 cm−1 , which corresponds to the center of the IR absorption band of the C=O stretching vibration of the
ureido group (R-HN-(C=O)-NH-R) in BAT. Both images display an area of 9 x 5 µm2
in a linear color scale in which black represents the minimal height and minimal scattered near-field intensity respectively. The line plots show profiles perpendicular to the
stripe pattern as indicated by the dashed lines in the images. In order to improve the
signal-to-noise ratio, the profiles are averaged over 20 single lines (gray points) and a
16-term moving average was applied (black line).
Topography as well as near-field image clearly reveal the line pattern of the laterally
111
Figure 10.6.: Simultaneously recorded topography (A) and near-field (B) images
(9 x 5 µm2 ) at the resonance frequency (1711 cm−1 ) of the ureido group of BAT. Black
represents the lowest features in the topography image and the minimal detected signal
in the near-field image. The line plots show 16 term-moving average profiles perpendicular to the stripe pattern averaged over 20 lines as indicated by the dashed lines in the
images.
structured SAMs. A contrast inversion among topography and near-field response is
observed when comparing both images (see also the corresponding line profiles). In the
topograph (Fig. 10.6 A) the narrow lines of BAT appear bright whereas they appear
dark in the near-field image (Fig. 10.6 B). This is due to the different contrast mechanisms of both techniques. While the topographic AFM contrast depends on the height
difference between both molecules the optical near-field response probes the dielectric
properties of the molecules. For ODT (total length: 2.5 nm) and BAT (total length:
3.7 nm) a theoretical height difference of 1.2 nm is expected which is in good agreement
with the measured height difference of ∼1.5 nm presented in the line profile. Because
of this height difference the BAT stripes appear brighter when looking at topography.
Concerning the near-field contrast the intensity of the scattered light is less for the BAT
regions caused by an attenuation of the near-field due to the dielectric properties (e.g.
absorption) of the biotin monolayer.
In order to investigate the spectral dependence of the near-field contrast measurements
were carried out at five specific wavenumbers: ν̃ = 1800, 1722, 1711, 1692 and 1640 cm−1 .
112
10.3.4. s-SNIM
Figure 10.7 displays the normalized IRRAS spectrum of BAT (solid gray line) in comparison to the normalized near-field contrast (black dots) in the spectral range from
1850 to 1600 wavenumbers. Additionally to the calculated near-field contrast the corresponding near-field images and one typical topograph are presented. For calculating the
near-field contrast first the near-field intensities of BAT and ODT were determined by a
histogram analysis of regions of interest at each laser frequency. From these intensities
the near-field contrast was calculated according to Equation 10.1 and normalized to 1.
C =1−
IBAT
IODT
(10.1)
C = near-field contrast, IODT (ν̃) = scattered near-field intensity of ODT,
IBAT (ν̃) = scattered near-field intensity of BAT
ODT was used as an internal reference because it has no absorption bands within the
investigated spectral region (Fig. 10.5).
Comparing the normalized IRRAS spectrum of BAT to the relative near-field contrast
reveals that the near-field contrast reflects the same frequency dependence as the far-field
absorption of the BAT SAM (Fig. 10.7). Although the IRRAS spectra of both SAMs
Figure 10.7.: Normalized IRRAS spectrum of BAT (solid gray line) and normalized
near-field contrast (black dots) with corresponding near-field images at five different
wavenumbers. A typical topograph is displayed at the top (gray).
113
Figure 10.8.: Simplified model of artifacts in near-field contrast due to a changing height
of the intermediate layer between tip and sample substrate. With increasing distance
between tip and gold surface the interaction between dipole and mirror-dipole becomes
attenuated.
show no far-field absorption at 1800 cm−1 a weak near-field contrast is observed. This
weak contrast is partly attributed to the different heights of the molecules (Fig. 10.8).
Due to the strong distance dependence of the scattered near-field signal with (z+a)3
(Equation 6.26) the thicker BAT layer causes a reduction of the signal with z/a of 3%.
However, any artificial contrast based on height differences between the SAMs should
be independent of wavelength. To this end these contributions can be separated by
measuring the image contrast over several wavelengths.
Topographical and optical lateral resolution were determined at one of the boundaries
between ODT and BAT (Fig. 10.9). The line profile was averaged over five rows and
an error function line was fitted to the data points in order to evaluate the step resolution. Edge width resolution between the 10% and 90% positions of the slope amounts
to ∼70 nm for topography and ∼90 nm for the near-field at a frequency of 1711 cm−1
(λ = 5.58 µm) which is comparable to a diffraction dependent resolution of approximately λ/60. Assuming a cross-sectional area of 21.4 Å2 per sulfur atom of an alkylthiolate [27] results in a molecular density of 7x10−24 mol/nm2 corresponding to 4 molecules
per nm2 . Consequently a detection area of approximately (90 nm)2 leads to a lower
detection limit of 5x10−20 mol corresponding to 3x104 molecules of BAT or 27 attogram.
Most likely the sensitivity is under estimated due to the disordering of the BAT-SAM
which was proved by the infrared spectra. The achieved sensitivity is consistent with
s-SNIM measurements on a tobacco mosaic virus performed by Brehm et al. [146].
114
10.3.4. s-SNIM
Figure 10.9.: Topography (A) and near-field (B) line plots of the slope of a stripe. An
error function line fit to the data points evaluates the step resolution. Five rows were
averaged to obtain the data points. The dotted lines mark the 90 and 10% positions
that were used to calculate the lateral resolution.
10.3.4.1. Theoretical predictions
Using a dipole-mirror dipole model as previously described in chapter 6.3.2 (page 61) it
could be shown in the following that the recorded intensity of the near-field spectrum
for thin film of a strong absorber, such as the urea group in biotin, is dominated by the
absorption cross-section. For this reason the near-field contrast shows an absorption-like
course instead of a dispersion-like spectrum as investigated by other groups.
In order to calculate the scattering and absorption cross-section of the SAMs their
complex dielectric constants are required. Since the complex dielectric constant is related
to the complex refractive index (see Equation 6.11) it can be calculated when imaginary
part (absorption) and real part (refractive index) of the complex refractive index are
known. The imaginary part can be obtained from the measured spectra of BAT and
ODT whereas the real part can be calculated using the Kramers-Kronig relation:
Z ∞
2
ν
0
n(ν ) = n∞ + P
k(ν) 2
dν
(10.2)
π
ν − ν 02
0
ν = frequency, P = principal value of the Cauchy integral, k = absorption coefficient
The refractive index of both thiolates is assumed to be 1.43 [8]. Following the dielectric constant is calculated using Equation 6.11. For the investigated CO region (18501600 cm−1 ) a maximal absorption (k) of 0.09 at 1711 cm−1 were obtained whereas the
corresponding value for the ODT-SAM is only 0.008. The refractive index variations
are 3% for BAT and 0.4% for ODT. Subsequently, the calculated dielectric constants
are used to estimate the effective polarizability of the dipole-sample system according
115
to Equation 6.26 with
β=
εSAM − εmedium
εSAM + εmedium
(10.3)
β = constant of proportionality, ε = dielectric constant
From the effective polarizability of the system the scattering and absorption crosssections can be calculated according to chapter 6.3.2 (page 61). The maximum absorption cross-section σabs results to 32 nm2 for BAT and 3 nm2 for ODT at 1711 cm−1
(for z=0). In contrast σsca does not exceed 0.03 nm2 for both SAMs in the whole investigate region from 1850 to 1600 cm−1 . Since the absorption cross-section is a factor 102
to 103 larger than the scattering cross-section the contribution of absorption dominates
the line shape of the near-field signal. Thus the spectral response of the near-field signal
is very similar to the far field absorption spectrum.
These findings are in agreement with recently published results from Aizupurua et al.
[115]. They investigated PMMA (poly(methylmetacrylate) layers of different thickness
on gold. With decreasing PMMA thicknesses the spectral features changed from a
dispersive-like to an absorption-like signature.
116
11. Detection of hybridization in DNA
monolayers by IRRAS, AFM and
s-SNIM
Contents
11.1. Introduction
. . . . . . . . . . . . . . . . . . . . . . . . . . . . 118
11.2.1. Oligonucleotides . . . . . . . . . . . . . . . . . . . . . . . . .
119
11.2.2. Sample preparation
. . . . . . . . . . . . . . . . . . . . . . .
119
11.2.2.1. Preparation of solutions . . . . . . . . . . . . . . . .
119
11.2.2.2. MCH and DNA SAM formation . . . . . . . . . . .
120
11.2.2.3. Hybridization of DNA SAMs . . . . . . . . . . . . .
120
11.2.2.4. Preparation of samples for nanografting . . . . . . .
121
11.2.2.5. Nanografting of MCH and ssDNA . . . . . . . . . .
121
11.2.3. Compressibility studies
. . . . . . . . . . . . . . . . . . . . .
124
11.2.4. Surface Plasmon Resonance (SPR) spectroscopy . . . . . . .
126
11.2.5. Ultraviolet (UV) spectroscopy . . . . . . . . . . . . . . . . . .
126
126
11.2.7. Calculated infrared spectra . . . . . . . . . . . . . . . . . . .
127
11.2.8. s-SNIM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
127
11.3. Characterization of DNA SAMs by IRRAS . . . . . . . . . . 128
11.3.1. Calculated infrared spectra of DNA . . . . . . . . . . . . . .
128
11.3.2. IRRAS experiments with MCH and DNA SAMs . . . . . . .
131
11.3.2.1. IRRAS on homogenous MCH SAMs . . . . . . . . .
131
11.3.2.2. Impact of backfilling with MCH on DNA SAM spectra133
11.3.2.3. Comparison between a MCH spectrum and ssDNA
and dsDNA spectra . . . . . . . . . . . . . . . . . .
133
11.3.2.4. IRRAS on DNA SAMs . . . . . . . . . . . . . . . .
134
117
Chapter 11. Detection of hybridization in DNA monolayers by IRRAS, AFM and s-SNIM
11.3.2.5. IRRAS study on ageing of DNA SAMs . . . . . . .
136
11.3.2.6. Impact of water on DNA SAM spectra . . . . . . .
138
11.4. Characterization of nanografted DNA and DNA SAMs using AFM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 139
11.4.1. Height studies on nanografted DNA structures . . . . . . . .
139
11.4.1.1. DNA heights before and after hybridization . . . . .
139
11.4.1.2. DNA heights after each fabrication step . . . . . . .
142
11.4.2. DNA compressibility study . . . . . . . . . . . . . . . . . . .
144
11.4.3. SPR study on DNA SAMs . . . . . . . . . . . . . . . . . . . .
144
11.4.4. Melting curves . . . . . . . . . . . . . . . . . . . . . . . . . .
148
11.4.5. Summarizing discussion on the height decrease of dsDNA upon
nanografting of ssDNA . . . . . . . . . . . . . . . . . . . . . .
150
11.5. Characterization of DNA nanostructures using s-SNIM . . 152
11.6. Summarizing conclusion . . . . . . . . . . . . . . . . . . . . . 157
11.1. Introduction
Thiol modified DNA molecules tend to form disordered films on gold surfaces. Besides
the chemisorption of the thiol group to the gold also the DNA bases especially the carbonyl and amino groups can interact with the gold surface. As outlined in the DNA
theory chapter backfilling with MCH is commonly used to suppress non-thiol interactions between gold and DNA. In order to gain experience with the formation of DNA
SAMs in IRRAS experiments the influence of MCH on DNA spectra was investigated.
Therefore DNA SAM formation with/without MCH backfilling and also the MCH incubation time was studied in detail. Brewer et al. reported that water (water bending
mode around 1638 cm−1 ) in DNA monolayers can mask the spectral region that contains most information about hybridization [166]. To this end the impact of water on the
DNA IRRAS spectra was analyzed. The experience gained in the pre-characterization
of DNA SAMs was used for preparing s-SNIM measurements with nanostructured DNA
monolayers. For direct comparison of SNIM contrast from different molecules it is desirable investigating molecules and reference molecules in close proximity to each other.
This allows imaging all structures within one step. A well-suited technique to fabricate
nanostructures in close proximity to each other is nanografting. In addition nanografting allows checking in situ the nanofabrication process using AFM height measurements.
After nanofabrication the samples were used to test the feasibility of DNA hybridization
detection by s-SNIM. Because of MCH molecules were chosen as spectral reference it
was important to know the infrared spectrum of MCH in comparison to ssDNA and
dsDNA spectra. Since the samples for s-SNIM were prepared at ELETTRA in Basovizza, Trieste (Italy) and transported to Bochum IRRAS experiments about the ageing
of DNA SAMs were necessary to check sample stability.
118
11.2.1. Oligonucleotides
HPLC purified single-stranded oligonucleotides were purchased from biomers.net GmbH
as dried aliquots with an amount of 4 or 10 nmol per aliquot. Sequences are listed
in Table 11.1. Probe DNA was modified at the 5´ terminus with a six carbon linker
containing a thiol group at the end. Complementary target DNA was used unmodified
or labeled with a biotin at its 5’-end. Duplex DNA was formed by hybridization of probe
and target strand.
name
HS-ssDNA
(probe)
comp ssDNA
(target)
ssDNABio
(target)
sequence
HS-C6 H12 -5´-AGA TCA GTG CGT CTG TAC TAG CAC A-3´
5´-TGT GCT AGT ACA GAC GCA CTG ATC T-3´
Biotin-5´-TGT GCT AGT ACA GAC GCA CTG ATC T-3´
dsDNA
(duplex)
3´-TCT AGT CAC GCA GAC ATG ATC GTG T-5´
dsDNABio
(duplex)
3´-TCT AGT CAC GCA GAC ATG ATC GTG T-5´-Biotin
Table 11.1.: Sequences of oligonucleotides used in the experimental part of this chapter.
Nucleobases participating in the secondary structure are typed in red.
Lowest energy secondary structures of single-stranded oligonucleotides predicted by
OligoAnalyzer3.1 (IDT) are shown in Figure 11.1. Probe and target strand have a
hairpin structure containing a hairpin-like stem of 4 base pairs, a 10 base loop, and a
6 base tail. These structures are considerable stable, having predicted ∆G◦ 20 values of 4.71 kcal/mol (target) and -4.01 kcal/mol (probe). Sequences containing intermolecular
hairpin structures can also form intermolecular self-dimers (target-target and probeprobe) with similar stability than the secondary structure. Concentration-dependent
thermal melting studies of probe and target ssDNA and van´t Hoff analysis indicate
that 90% of probe strands and 100% of target strands exist as hairpins, rather than
dimers at 20◦ C and 1 µM ssDNA in STE-buffer [35].
11.2.2.1. Preparation of solutions
All solutions were prepared using Milli-Q or HPLC water (J.T.Baker) (18 M Ohm cm−1
resistance). STE buffer consists of 1 M NaCl in TE buffer (10 mM Tris buffer pH 7.2
and 1 mM EDTA, Sigma). 6-Mercapto-1-hexanol (MCH, ≥97%, Fluka) was prepared
119
Figure 11.1.: Lowest energy secondary structures as predicted by OligoAnalyzer3.1
(IDT) under conditions of 1 µM strand concentration and 1 M NaCl. Probe strand
as well as target strand contain a 10 base loop with a 4 base pair stem.
as a 1 mM solution in STE buffer. Buffer solutions were filtered prior to use through a
0.22 µm pore size filter. Ethanol was used in p.a. quality. Urea was employed as 6 M
solution in pure water.
11.2.2.2. MCH and DNA SAM formation
As gold substrates either sputter deposited gold (Anfatec Instruments AG) or template
stripped gold was utilized. Sputter deposited gold substrates were prior to use cleaned
with hot piranha solution (7:3 mixture of H2 SO4 and H2 O2 ) for approximately 10 min,
rinsed thoroughly with HPLC water, and temporary stored in HPLC water (detailed
description see section 7.1.1 (page 76)). Template stripped gold (TSG) was prepared
as described in section 7.1.2 (page 76). Stripping precursors were opened immediately
before use. MCH SAMs were grown by exposing gold substrates into a 1 mM ethanolic
or STE-buffer solution for 30 or 90 min. Low density DNA SAMs were prepared by immersing a gold surface for 1.5 min unless mentioned otherwise into a solution containing
1 µM HS-ssDNA. Subsequently the surface with the DNA monolayer was back-filled by
exposure to a solution containing 1 mM MCH in STE buffer for 30 min unless mentioned
otherwise.
11.2.2.3. Hybridization of DNA SAMs
Double-stranded DNA was obtained after incubation of immobilized HS-ssDNA in 1 µM
complementary ssDNA (target) in STE buffer at room temperature for a least 6 hours.
120
11.2.2.4. Preparation of samples for nanografting
Nanografting experiments were carried out on TSG surfaces. Immediately after stripping
the freshly exposed gold surface was immersed into a solution containing the desired thiolated molecule. Depending on the DNA nanografting experiment different background
SAMs were used. Preparation of different background SAMs is listed in Table 11.2.
After washing, the sample was dried in a stream of nitrogen and glued onto a steel plate
SAM
DNA SAM
MCH SAM
solution
1. step: 1 µM HS-ssDNA
in STE buffer
2. step: 1 mM MCH in STE buffer
3. step: thoroughly washing with 10 mM NaCl
1. step: 1 mM MCH in STE buffer or EtOH
2. step: thoroughly washing with water and/or EtOH
incubation time
1.5 min
30 min
over night
Table 11.2.: Preparation of different background SAMs for DNA nanografting
(AFM disc, d=15 mm, Plano GmbH) using a very small drop of PDMS. In order to cure
the PDMS the substrate was put on a prewarmed heater plate (approx. 100◦ C) for less
than 10 s (to avoid damages of the SAM). Subsequently it was cooled down for at least
1 minute on a metallic surface with good heat conductivity. This process was repeated
until the PDMS was fully cured (1-3 times). Little amount of PDMS as well as mixing
it some days before usage and storing it in a fridge accelerates curing. Spreading the
PDMS under the substrate facilitates curing too and prevents tilting of the gold surface.
11.2.2.5. Nanografting of MCH and ssDNA
All nanografting experiments were carried out with a conventional Solver Pro AFM
(NT-MDT) with a closed liquid cell in contact mode.
The steel plate with the fixed sample substrate was mounted on the AFM sample holder
and a worn out cantilever (e.g. with a blunted tip) was used to scratch a sign into the
SAM covered gold surface. This sign allows aligning the cantilever optically to an origin
which facilitates finding back the nanografted pattern easily. For nanografting ”hard”
NSC36C cantilevers (MikroMasch, nominal force constant 0.6 N/m, tip radius <10 nm)
were employed. To this end surplus cantilevers A and B were carefully removed by locally
applying mechanical force prior to cantilever installation. After mounting the marked
gold substrate in a closed liquid cell filled with 850 µl of STE buffer the AFM laser spot
was aligned onto the cantilever. Then the sample mark was roughly positioned under the
cantilever using the x-y-translation stage of the sample holder. Thereby one has to pay
attention to not apply to much tension onto the accordion like membrane of the liquid
cell otherwise the sample drifts during scanning. Pre-alignment should be performed
in the same solvent that is later on used for nanografting since the alignment depends
121
Figure 11.2.: Schematic step by step description for the nanografting process.
122
Figure 11.3.: Topographs were measured in TE-buffer in contact mode with a low loading
force (<1 nN)
.
123
on the refractive index of the solvent. In order to avoid unnecessary displacement of
SAM molecules by thiols from solution exposure to nanografting solution should be
kept as short as possible. For this reasons pre-alignment was carried out in STE buffer.
Afterwards the liquid cell was removed from the AFM and the solvent was replaced with
nanografting solution containing either 0.1 mM MCH in STE buffer or 10 µM HS-ssDNA
in 1:1 ethanol/STE buffer solution (v/v). During exchange of solution it is important not
to move the substrate within the liquid cell otherwise a new sample-tip alignment might
be necessary. After remounting the liquid cell a fine alignment was done and the SAM
was imaged under low load to select a well-suited area for nanografting. Subsequently
loading force was increased to typically 50 to 80 nN and an area with the desired pattern
dimensions was scanned. During scanning the AFM tip removes locally molecules from
the SAM and thiols from the surrounding solution chemisorb onto the freshly exposed
gold area following the scanning track of the AFM tip. In order to check if the grafting
was successful and to select the next grafting position the scan area size was increased
and scanned with a low loading force to prevent damaging the nanografted structure.
Immediately after finishing the nanografting process the sample in the liquid cell was
thoroughly washed with STE buffer. Figure 11.2 displays the nanografting process step
by step.
Imaging of the nanografted structures was performed in STE buffer using a ”soft”
CSC38B cantilever (MikroMasch, nominal force constant 0.03 N/m, tip radius <10 nm)
at low loading forces (typically <1 nN).
Image processing
Image processing was performed using Scanning Probe Image Processor (SPIP, Image
Metrology). AFM images were corrected using a linear or parabolic flatten function.
Heights were determined either by line profiling using SPIP or by averaging over regions
of interest using Mathematica 5.2 (Wolfram Research).
All measurements were performed in STE-buffer with a ”soft” CSC38B cantilever (MikroMasch, nominal force constant 0.03 N/m, tip radius <10 nm). Initially the force constant
of the cantilever used for compressibility studies was determined. To this end the minimal, typical and maximal force constants from the cantilever data sheet were plotted
against the corresponding resonance frequencies and fitted to a quadratical equation
(Fig. 11.4 A). Using this equation the force constant of the cantilever can be estimated inserting its actual resonance frequency which can be detected with the AFM
(Fig. 11.4 B).
Prior to measurements the force out of contact was set to zero by moving the reflected
beam from the cantilever into the middle of the 4-segment photodiode or by defining
a certain current value as offset. Then a force-distance curve (FZ curve) was recorded
(Fig. 11.4 C). Dividing the force constant by the slope of the force-distance curve yields
the sensitivity of the AFM set-up (nN/nA) which allows to calculate the loading force
from the applied set point or vice versa.
124
For compressibility measurements the sample was scanned with different loading forces
ranging from 0.3 up to 10.5 nN (0.3, 0.75, 1.2, 1.5, 2.89, 4.5, 7.5, and 10.5 nN). Reversibility of the compression is studied by setting the loading force back to its initial
value of 0.3 nN.
Figure 11.4.: Example how to determine the force constant of a cantilever and the
sensitivity of an AFM set-up. From the sensitivity the required set-point for a certain
loading force can be calculated.
Image processing
Image processing was performed using Scanning Probe Image Processor (SPIP, Image
Metrology). AFM images were corrected using a linear or parabolic flatten function.
Heights were determined by averaging over regions of interest using Mathematica 5.2
(Wolfram Research).
125
11.2.4. Surface Plasmon Resonance (SPR) spectroscopy
SPR measurements were carried out by Dr. Ch. Grunwald. Data are presented with
his permission. Thin glass slides (D263, Schott) were coated with a primer layer of 12
Å titanium and followed by the deposition of 475 Å gold. For preparation a Leybold
Inficon XTC/2 metal evaporator was used. The coating was done at room temperature and a pressure of approximately 10−7 bar. DNA SAM formation was carried out
as described in section 11.2.2.2 (page 120). Prepared DNA SAMs were washed with
STE-buffer and dried in a stream of nitrogen. SPR experiments were carried out in a
SR7000DC Dual Channel spectrometer (Reichert). Prior to measurements the installed
sample was equilibrated in STE-buffer at a constant flow rate of 20 µl/min over night.
Experiments were performed at RT in STE-buffer solution at a flow rate of 20 µl/min.
DNA SAM hybridization and subsequent washing with a 1:1 mixture of ethanol/STEbuffer (v/v), pure water and 6 M urea was accomplished by sequential injections of
250 µl of each substance over a time period of 12.5 min. Between each injection the
sample was equilibrated in STE-buffer solution for 21 min.
11.2.5. Ultraviolet (UV) spectroscopy
Duplex DNA was prepared by mixing equal amounts of HS-ssDNA and complementary
ssDNA in STE-buffer over night at room temperature. Prior to thermal melting DNA
solution was diluted 1-fold with either pure ethanol or pure water yielding a total DNA
concentration of 250 nM. Melting experiments were performed using a Jasco V-630
spectrometer equipped with a temperature-controlled sample holder. For experiments
in aqueous STE solution the absorbance at 260 nm was recorded from 20 to 100◦ C.
During experiments with EtOH/STE buffer solution the temperature interval was set
from 20 to 80◦ C. The heating/cooling rate was 1◦ C/min.
SAMs for IRRAS measurements were prepared on sputter deposited gold. IRRAS spectra were recorded on a Vertex V80 (Bruker) spectrometer equipped with a globar source,
a KBr beam splitter, a BaF2 polarizer (LOT), a grazing incident reflection unit and a
liquid nitrogen cooled MCT detector. The gold grid polarizer was used to obtain ppolarized radiation before reflecting off the sample at an incident angle of 80◦ from the
surface normal. Spectra were recorded at room temperature with a resolution of 4 cm−1
and a coadding of 256 spectra. Fourier transformation of all spectra was carried out
with Mertz phase correction and triangular apodization. Evacuation time for the sample chamber was 5 min. As reference either a bare gold substrate or a MCH SAM was
used.
For evacuation time-dependent spectra reference as well as sample were evacuated for 1
hour in total and spectra were recorded after 5, 10, 20, 30, 40, 50, and 60 minutes of
evacuation.
126
11.2.7. Calculated infrared spectra
Spectra processing
Spectra were evaluated using OPUS-Software 6.5 (Bruker) region from 950 to 3000 cm−1 .
Difference spectra were calculated by subtracting ssDNA or dsDNABio spectra, respectively, from dsDNA spectra.
11.2.7. Calculated infrared spectra
Theoretical infrared spectra were based on density functional theory (DFT) calculations
of the individual DNA bases and complementary base pairs from Brewer et al. [166].
Calculated spectra for the DNA sequences used in this work were obtained by using
weighted averages of the Brewer data.
11.2.8. s-SNIM
SNIM measurements were carried out as described in chapter 9 (page 91). Each image
was recorded at a scan rate of 0.5 Hz and time constant of 1 ms at the lock-in amplifier.
Samples for s-SNIM were prepared at ELETTRA in Basovizza, Trieste (Italy) and transported under ambient conditions to Bochum, Germany. With exception of the transport
samples were stored in a nitrogen atmosphere inside an exsiccator. Preparation of the
nanografted structures is described in section 11.2.2.5 (page 121).
Image Processing
Image processing was performed using WSxM imaging software 4.0 Develop 10.4 (Nanotec Electronica) [167]. Raw images were corrected using a linear flatten function and
for presentation the image contrast was enhanced.
Near-field data were evaluated using Mathematica 5.2 (Wolfram Research).
127
11.3. Characterization of DNA SAMs by IRRAS
The infrared spectrum of DNA has many absorption bands in the spectral region from
500 to 2000 wavenumbers. Their intensity and position depends mainly on the geometry
of the nucleic acid. Typical frequency regions of the vibrational bands are:
500−1000 cm−1 : sugar-phosphate-backbone vibrations
1000−1300 cm−1 : phosphate vibrations
1300−1600 cm−1 : deformation vibrations of the bases coupled to the sugar
vibration by the glycosidic linkage
−1
1600−1800 cm : vibration modes of the double bonds of the bases.
Position depends on base pair packing
The frequency region from 1800 to 1500 cm−1 is of special interest for studying hybridization in DNA by infrared spectroscopy.
11.3.1. Calculated infrared spectra of DNA
Figure 11.5 shows DFT calculated spectra of the individual bases and the complementary base pairs in the frequency range from 1800 to 1575 cm−1 according to Brewer et al.
[166]. Spectra for the individual bases are displayed in the upper part of diagram 11.5.
Dashed lines correspond to the heterocyclic purine bases (A and G) and dotted lines
to the homocyclic pyrimidine bases (T and C). Complementary bases have the same
color (A, T = red; C, G = blue). A peak assignment is given in table 11.3. Vibrational
modes below 1680 cm−1 are primarily due to NH2 and N-H bands and ring deformations
whereas modes around 1700 cm−1 are assigned to carbonyl stretching vibrations. The
lower part of diagram 11.5 shows calculated spectra for the complementary base pairs.
Upon base pair formation vibrational modes of N-H and NH2 are blue shifted to higher
frequencies and carbonyl bands are red shifted to lower frequencies. The shifts are due
to hydrogen bonding which constrain amine vibrations and weaken the force constant
of the carbonyl groups.
Figure 11.6 depicts calculated infrared spectra in the spectral range from 1780 to
1580 cm−1 of ssDNA and dsDNA for the DNA sequence employed in this work. Comparing the spectra of random coil ssDNA (dotted gray line) and 100% hybridized DNA
(dotted black line) the main difference is the rise of an infrared mode around 1675 cm−1
upon hybridization. Additionally the other modes are altered in shape, center frequency
and intensity. The vibrational mode around 1750 cm−1 decreases, at 1720 cm−1 the
peak rises and around 1640 cm−1 the peak intensity increases slightly. According to
studies of Georgiadis and coworkers, who employed the same DNA sequence as used in
the experiments in this chapter, 90% of this ssDNA sequence forms a hairpin structure
at a concentration of 1 µM in STE-buffer [35]. Because secondary structure is based
on intramolecular base pair formation it influences the spectral response following the
128
11.3.1. Calculated infrared spectra of DNA
Figure 11.5.: Calculated infrared spectra for the individual bases and complementary
base pairs according to Brewer et al. [166]. Peak assignment is given in table 11.3.
Figure 11.6.: Calculated infrared spectra for the DNA sequence used in this thesis.
129
base, basepair
adenine (A)
thymine (T)
guanine (G)
cytosine (C)
AT
GC
center peak
group
−1
[cm ]
1583
NH2 bending
C=N stretching (ring
1630
NH2 bending
ring deformation
1649
C−C stretching (ring
1705
C=O stretching
1755
C=O stretching
1580
NH2 bending
ring deformation
1630
NH2 bending
ring deformation
1742
C=O stretching
1598
NH2 bending
1655
NH2 bending
1734
C=O stretching
1605
NH2 bending
ring deformation
1656
NH2 bending
1676
N−H bending
C=O stretching
1754
C=O stretching
1629
NH2 bending
N−H bending
1677
NH2 bending
C=O stretching
1717
N−H bending
C=O stretching
deformation)
deformation)
deformation)
deformation)
deformation)
Table 11.3.: Peak assignment for the calculated spectra of DNA bases and base pairs in
the spectral region from 1785 to 1575 cm−1 according to [166, 168]
130
11.3.2. IRRAS experiments with MCH and DNA SAMs
same trend as hybridization does. Therefore the secondary structure has to be considered in theoretical prediction of the DNA spectra. The calculated spectrum for 90%
hairpin structure and 10% random coil ssDNA is displayed as solid gray line. Furthermore Gao et al. reported a hybridization efficiency of only 50% for this hairpin forming
DNA sequence [35]. For this reason also a spectrum for a hybridization efficiency of
50% is shown as solid black line. Comparing the hairpin spectrum (solid gray line) and
the partly hybridized DNA spectrum (solid black line) main differences are observed at
1720 cm−1 and 1680 cm−1 .
The calculated infrared spectra illustrate that the infrared spectrum of DNA in the
spectral range between 1800 to 1500 wavenumbers strongly depends on DNA secondary
structure and degree of hybridization. When comparing the experimental and the calculated spectra one has to keep in mind that also hydration may affect the vibrational
modes and the calculations are based on gas phase predictions. In addition Brewer et
al. used only DNA bases for cluster calculations neglecting influences from the sugarphosphate backbone of the DNA.
11.3.2.1. IRRAS on homogenous MCH SAMs
Figure 11.7 presents IRRAS spectra of homogeneous MCH SAMs on gold immobilized
from aqueous STE-buffer solution (red) and ethanolic solution (black). Peak assignment
is given in table 11.4. When immobilized from ethanolic solution (black spectrum)
characteristic absorptions features of MCH appear. The local frequencies of the CH2
vibrational modes (νs at 2854 cm−1 and νa at 2925 cm−1 ) confirm a disordered liquid-
Figure 11.7.: IRRAS spectra of MCH immobilized on gold from an aqueous STE-buffer
solution (red line) and from ethanolic solution (black line). Peak assignment is given in
table 11.4.
131
[cm−1 ]
950-1070
1070
1150-1350
1265
1350-1480
1462
2854
2925
2960
3200-3600
vibrational mode
chain rocking and twisting modes
C−O stretch
chain wags and twists
CH2 bending
CH2 scissor def.
CH2 sym. stretch
CH2 antisym. stretch
CH3 antisym. stretch
O−H stretch
Table 11.4.: Peak assignment for the IRRAS spectrum of MCH in the spectral region
from 3600 to 950 wavenumbers according to references [169]
like conformation of the molecules which is characteristic for short-chain alkylthiols [29].
However, SAM growth from STE-buffer show only very weak absorption peaks. Low
intensities of methylene stretching modes in the high frequency region and the C-O
stretching mode at 1070 cm−1 indicate low coverage and high disorder of the SAM
immobilized from STE-buffer. Even immobilization times of 90 min did not improve
SAM growth from aqueous solution.
Choice of solvent is an important but poorly understood parameter in SAM growth
from solution which influences kinetics and mechanism of assembly. Studies suggest
that SAM formation of alkanethiols from nonpolar hydrocarbon solvents improve the
kinetics of formation in some cases (e.g. heptane [170] or hexane [171]) but may result
in less organized SAMs compared to the organization of SAMs immobilized from ethanol.
Less studies report on SAM growth from aqueous solutions, which are poor solvents for
alkanethiols. Nevertheless SAMs can form from aqueous solution containing micelles of
ionic or nonionic surfactants which form around alkylthiols and aid their diffusion to the
gold surface. SAMs grown in such solvents seem to promote densely packed monolayers
[172]. At present, we do not understand the differences in SAM formation in ethanol and
in buffer. Possible explanations might include different conformations of the adsorbed
MCH molecules (striped and cluster-like SAM structures) [173].
Spectra obtained from buffer solutions exhibit a small mode around 1714 cm−1 that can
not be assigned to the vibrations of the chemical groups within the MCH molecule. This
peak might be from physisorbed EDTA which is a buffer ingredient and contains four
carboxyl groups.
132
11.3.2.2. Impact of backfilling with MCH on DNA SAM spectra
Although MCH immobilized from STE-buffer shows only weak spectral features backfilling with MCH strongly affects DNA molecular orientation. Figure 11.8 A displays
spectra of a high density DNA SAM (immobilization over night) with (red line) and
without (black line) subsequent MCH treatment. After backfilling with MCH intensities of DNA vibrational modes are dramatically increased. This is most likely due to
displacement/removement of unspecific DNA strand interactions with the gold surface
by MCH molecules causing the DNA molecules to rearrange in a more upright manner
in which only the sulfur group is attached to the gold. Such a reorientation is consistent
with findings of Tarlov and coworkers who observed a better accessibility of the DNA
strands for hybridization after MCH treatment [39, 40].
Graph 11.8 B presents the impact of incubation time in MCH solution on the DNA
spectra. For an extended incubation time of 90 min no significant change in spectral
response was observed compared to a DNA SAM post-treated for only 30 min with
MCH. Therefore it is deduced that most unspecific interactions between DNA and gold
surface are removed within 30 min.
Figure 11.8.: Influence of MCH backfilling on IRRAS spectra. A) IRRAS spectra of
high density DNA SAMs backfilled with MCH (red spectrum) and without MCH (black
spectrum). B) IRRAS spectra of DNA SAMs after treatment with MCH for 30 min (red
spectrum) and 90 min (black spectrum).
11.3.2.3. Comparison between a MCH spectrum and ssDNA and dsDNA spectra
Figure 11.9 shows the spectrum of a MCH SAM immobilized from STE-buffer (red line)
in comparison to a low density DNA/MCH SAM before (gray line) and after hybridization (black line). All spectra were measured versus a bare gold reference. Peak intensities
of MCH are small compared to DNA modes (< 20%). In the following MCH SAMs from
STE-buffer solution were used as reference samples for DNA IRRAS measurements for
two reasons: (i) with regard to the fast contamination of bare gold SAMs offer a more
defined reference surface. (ii) absorptive influences from the MCH molecules beneath
133
Figure 11.9.: IRRAS spectra of MCH immobilized from STE-buffer solution (red line)
in comparison to a mixed low density ssDNA/MCH SAM (1.5 min ssDNA SAM) before
(gray line) and after hybridization (black line) measured versus a bare gold reference
the DNA SAM and the hexyl linker of DNA molecules can be minimized using a MCH
SAM as reference. In addition also the effect of physisorbed EDTA is reduced because
sample as well as reference are prepared from buffer solution. When comparing the spectra of DNA before and after hybridization the last point is not that important because
both spectra should be influenced in the same manner. But considering comparability
between different data series it is important to have a defined reference sample.
11.3.2.4. IRRAS on DNA SAMs
Less is reported on hybridization detection of immobilized DNA using infrared spectroscopy [166, 174]. As mentioned before the most sensitive spectral region to detect
formation of double-stranded DNA is between 1800 and 1550 cm−1 due to the vibrational modes of the DNA bases. Upon hybridization the spectral response is changed
due to formation of hydrogen bonds between the DNA bases resulting in an alteration
of vibrational excitation energies and vibrational properties of the chemical groups.
Figure 11.10 presents IRRAS spectra of a low density DNA SAM (1.5 min immobilization) before (red spectrum) and after (black spectrum) hybridization and the corresponding difference spectrum (lower diagram). Peak assignment is given in table 11.5.
Both DNA spectra exhibit similar spectral profiles. Upon hybridization the intensity
of the phosphate bands at around 1100 cm−1 and 1250 cm−1 increase due to a rising
number of phosphate groups from the complementary DNA strand and orientational
134
Figure 11.10.: IRRAS spectra for immobilized DNA before (red spectrum) and after
hybridization (black spectrum) with complementary DNA. The lower spectrum shows
the difference spectrum of the upper ones. Insets show the region from 1775-1575 cm−1
enlarged.
changes within the DNA SAM. Even vibrational modes within the spectral region from
1550 to 1750 cm−1 change in intensity and shape. As mentioned before this range is
of special interest for the detection of hybridization because of its changing spectral
response upon base pair formation. In addition a small peak at 1665 cm−1 gains intensity upon hybridization and another around 1710 cm−1 changes its peak shape. These
findings are in good agreement with the theoretical predicted spectral changes upon
hybridization (Fig. 11.6). However, the weak peak at 1665 cm−1 might indicate an even
lower hybridization efficiency than 50%.
135
peak center
position
[cm−1 ]
950-1070
1053
1070
1082
1150-1350
1238
1419, 1464
vibrational mode
2859
chain rocking
C-O-P vibrations
C-O stretching
PO2 − sym.
chain wags, chain twists
PO2 − antisym.
CH2 scissor deformation
CH2 bending
ring deformations
C=O
C=N stretching
exocyclic −NH2 bending
CH2 sym.
2888
2935
CH3 sym.
CH2 antisym.
2956
CH3 antisym.
1300 - 1550
1550 - 1750
group
C6 H12 chain (MCH, DNA linker)
DNA sugar-phosphate backbone
MCH
DNA phosphodiester backbone
DNA phosphodiester backbone
DNA bases (couple to glycosidic bond)
DNA bases
DNA bases
DNA bases
C6 H12 chain (MCH, DNA linker),
deoxyribose
thymine base
C6 H12 chain (MCH, DNA linker),
deoxyribose
thymine base
Table 11.5.: Peak assignment for the IRRAS spectra of immobilized DNA in the spectral
region from 3000 to 950 wavenumbers [169, 166, 174]
In order to prove that the spectral changes caused by hybridization figure 11.11 shows
spectra of a low density SAM after hybridization with unlabeled (black spectrum) and
biotinylated (blue spectrum) complementary ssDNA. The red spectrum displays the
difference both. The difference spectrum is similar to the spectrum of a biotinylated
alkylthiol SAM indicating the existence of biotinylated complementary DNA in the
hybridized DNA SAM. Differences might be caused due to varying orientation of the
biotin group within the two SAMs.
11.3.2.5. IRRAS study on ageing of DNA SAMs
Since the samples for s-SNIM measurements were prepared in Italy and transported to
Bochum within less than two weeks the influence of ageing on the spectral response
of DNA monolayers were studied on 18-days old monolayers. The DNA SAMs were
136
Figure 11.11.: IRRAS spectra of dsDNA hybridized with biotinylated complementary
DNA (red spectrum) and unlabeled complementary DNA (black spectrum). The difference spectrum of both is displayed below in blue. For comparability an IRRAS spectrum
of a biotinylated alkylthiol SAM is displayed.
Figure 11.12.: IRRAS spectra of fresh prepared DNA SAMs before and after hybridization (solid lines). Dashed spectra show the same SAMs after 18-days.
137
measured immediately after preparation, subsequently stored in an exsiccator under
inert atmosphere (nitrogen) and measured again after 18 days. No drastic changes were
observed. Small fluctuations might also result from slightly different positioning of the
sample in the spectrometer.
11.3.2.6. Impact of water on DNA SAM spectra
Brewer et al. [166] reported that the hybridization peak around 1655 cm−1 could be
masked by the strong scissoring peak of water around 1640 cm−1 . In order to exclude
such effects spectral response of DNA SAMs in dependence on the spectrometer evacuation time were investigated. Figure 11.13 depicts spectra of a low density ssDNA
SAM measured against bare gold in dependence of the sample chamber evacuation time
(5, 10, 20, 30, 40, 50, 60 min). A clear decrease in peak intensity for the band around
1430 cm−1 is shown and even the antisymmetric phosphate band at 1238 cm−1 is slightly
decreasing whereas no change is observed for the other spectral features. Especially not
for the region around 1650 cm−1 .
Figure 11.13.: Alteration of a ssDNA as function of spectrometer evacuation time.
138
11.4. Characterization of nanografted DNA and DNA SAMs using AFM
11.4. Characterization of nanografted DNA and DNA
SAMs using AFM
11.4.1. Height studies on nanografted DNA structures
In recent AFM studies height measurements with respect to reference areas such as the
underlying gold or ”reference” molecules were used to gain information about sample
surfaces and to characterize it. When working in liquids different solvents or reactions
with other dissolved molecules can cause height changes of immobilized molecules.
In the following all height referencing is done with respect to the gold surface (0 nm). To
this end MCH areas were used as reference. Since MCH does not change height during
DNA structure fabrication (i.e. DNA nanografting, hybridization and imaging) its height
was set to 1.2 nm [175] with respect to the gold surface. If not mentioned explicitly all
height measurements were carried out with low loading forces (typically <1 nN) in
contact mode in liquid environment.
11.4.1.1. DNA heights before and after hybridization
Figure 11.14 depicts topographs of nanostructured DNA before (A, E) and after hybridization (B, E) and corresponding line profiles (C,F). Measurements were carried
out in STE-buffer solution using contact mode with low loading forces (typically <1
nN). In order to improve signal-to-noise ratio line profiles were averaged over almost
the entire length of the stripes and the width of the squares. Small black features in
the topography are defects in the gold surface. In image A some bright spots appeared
after nanografting at the upper edge of the DNA stripe (bright stripe, upper part of the
topograph). These ”artifacts” are set to the mean height to improve image contrast.
After hybridization those bright spots disappeared. Most likely they are originated from
molecules shoved aside during nanografting (e.g. DNA and MCH aggregates) which
disassembled during hybridization in STE-buffer solution. In order to simplify matters
the discussion refers first only to the stripes (Fig. 11.14 A, B and diagram C). Later the
discussion is expanded to the patch-in-a-patch structure (Fig. 11.14 D, E and diagram F).
Image A and B show nanografted stripes of MCH (dark stripe) and DNA (bright stripe)
within a low density DNA SAM. Line profiles perpendicular to the stripes are plotted
in diagram C. Before hybridization the low density DNA SAM extends about 4.25 nm
from the gold surface whereas the nanografted DNA (DNA stripe) shows a height of
about 5.75 nm although both consist of the same DNA sequence. In a previous study
Liu and coworkers reported on two different surface reaction pathways of thiol selfassembly on gold: (i) unconstrained self-assembly in natural growth and (ii) spatially
confined self assembly (SCSA) in nanografting [65]. Due to the short persistence length
of only ∼1 nm of ssDNA unconstrained self-assembly results in a surface tethered coiled
and disordered conformation (Fig. 11.15) [39, 175]. Additionally the negatively charged
DNA phosphate-sugar backbones prevent dense packing because of electrostatic repul-
139
Figure 11.14.: Topographs of nanografted patterns of MCH (dark stripe and squares)
and DNA (bright stripe and squares) in a low density DNA SAM before (A, D) and after
hybridization (B, E) and corresponding line profiles perpendicular to the nanografted
stripes (C) and horizontal to the patch-in-a-patch structure (F). Small black features in
the topographs are defects in the gold surface.
sion among the strands. However, SCSA was found to improve density and molecular
ordering of the nanografted species. During nanografting the freshly exposed gold area
is spatially confined forcing the DNA molecules to assemble in an almost fully stretched
and standing up conformation [45, 79]. In order to quantitatively analyze the measured
heights the theoretical predicted fully stretched length of the used ssDNA is required.
Due to the secondary structure of the employed HS-ssDNA sequence an exact length
prediction is difficult and two assumptions have to be made: (i) the base paired part
forms a B-DNA conformation and (ii) the outer part of the hairpin loop is assumed
to have the length of 5 single-stranded bases. In both cases the calculated length is
overestimated. Steric constraints and electrostatic repulsion most likely result in shorter
conformations especially with regard to the length of the outer part of the loop. Con-
Figure 11.15.: Arrangement of ssDNA molecules after unconstrained assembly from
solution and SCSA during nanografting
140
Figure 11.16.: Theoretical predicted length of a 25-mer hairpin structured HS-ssDNA
and a 25-mer dsDNA
sidering the two assumptions, a base length of 0.4 nm for ssDNA [45], 0.34 for B-DNA,
and 1.2 nm for the hexane-thiol linker yields a maximum length of 6.96 nm for the
employed HS-ssDNA (Fig. 11.16). Comparing the measured height of the nanografted
ssDNA with the calculated length shows that the molecules are almost fully stretched.
The slight variation of about 1.21 nm with respect to the predicted length is most likely
due to imperfections in packing and/or the overestimated length.
After incubation in complementary ssDNA solution the height of immobilized DNASAM as well as that of the nanografted molecules is significantly increased (Fig. 11.14 C,
gray). This rise in height is characteristic for a reorientation of the DNA molecules upon
hybridization. For quantitative analysis of the measured DNA heights again a theoretical predicted length is required. Considering a base length of 0.34 for double-stranded
B-DNA, which is the most common dsDNA form in physiological conditions, yields an
expected maximum length of 9.7 nm for the employed 25-mer (including the hexane thiol
linker, Fig. 11.16). Comparing the predicted length with the height of the nanografted
DNA stripe, that extends about 8.75 nm from the surface, indicates an almost fully upright conformation of the hybridized molecules. However, the height of the DNA SAM
after hybridization is only about 6.75 nm. This is most likely due to the low probe
density providing only low lateral support for the hybridized DNA molecules.
Topographs D and E in Figure 11.14 show a patch-in-a-patch pattern, i.e. DNA (bright
0.5 x 0.5 µm2 squares) was nanografted in a before fabricated MCH patch (dark 1.5 x
1.5 µm2 squares). Diagram F presents the corresponding line profiles before (light gray)
and after hybridization (dark gray). The patch height before and after hybridization
follow the same trend as discussed above for the DNA stripes.
When comparing the heights of the DNA patch (5 nm) with that of the DNA stripe
(5.75 nm) before hybridization, the molecules within the stripe extend about 0.75 nm
more from the surface. In nanografting the probe density and consequently also the
141
height of the fabricated structures depends on the line density that is chosen during
nanografting [78, 79] (for a definition of the term ”line density” see chapter 3.2.1.1
(page 29)). Mirmomtaz et al. found that grafting over the same area more than once
increases packing density and accordingly height of the nanografted DNA structures [79].
Nevertheless, both DNA structures (patch and stripe) were written with a line density
of 1.28. This indicates that beyond the line density even the matrix into which the
molecules are grafted plays an important role. While the DNA stripe was nanografted
into a low density DNA SAM backfilled with MCH, the DNA patch was written into a
nanografted and consequently densely packed MCH matrix. Due to the low density and
high disorder a more precise and better replacement of the DNA/MCH SAM molecules
is expected during nanografting leading to a better ordering and higher packing density
within the nanografted structure.
After hybridization the height of the DNA patch increases only to about 7 nm. In
comparison to the height increase of the DNA stripe upon hybridization (8.75 nm)
this indicates less ordering and more entanglement of the strands in the ssDNA patch.
Mirmomtaz et al. have recently proven that less entanglement of ssDNA molecules
results in higher hybridization efficiencies [79].
11.4.1.2. DNA heights after each fabrication step
Figure 11.17 A displays the heights of nanografted DNA structures and the DNA SAM
after each fabrication step. Topograph 11.17 B shows the DNA structures after fabrication imaged in STE-buffer and scheme C assigns the structures to diagram A. A detailed
description of the fabrication steps is given in chapter 11 (page 117) and is depicted in
Figure 11.2 (page 122). Depending on the fabrication conditions height measurements
were carried out either in STE-buffer solution or an 1:1 ethanol/STE-buffer mixture
(v/v) using contact mode with low loading forces (typically <1 nN). Heights were determined by averaging over regions of interest. The standard deviation is about 1 nm
for all measurements.
All DNA structures show similar trends in height alteration during different fabrication
steps although the individual heights differ from each other. Most striking is the height
increase upon hybridization (dark grey bar) which was already discussed in the previous subsection. When comparing the measurements carried out in EtOH/STE-buffer
(black and light grey bars) with measurements in STE-buffer (all other bars) a solvent
dependent height change is detected. In all cases the height in EtOH/STE-buffer is
lower than in STE-buffer. This is most likely attributed to dehydration of the DNA by
the ethanol. The lower dielectric constant of ethanol in comparison to aqueous buffer
(at 25◦ C: DH2 O = 80.1, DEtOH = 24.3) causes the positive sodium-ions from the buffer
solution to replace water molecules from the negative phosphate backbone. This results
in a lower hydrophilicity of the DNA and most likely to a contraction of the molecules.
Interesting is that the height of the nanografted DNA seems to be more effected than
the height of the DNA SAM. For the SAM molecules height alters only about ∼0.5 nm
when changing the medium whereas the height of the nanografted DNA changes about
1.5 to 2 nm. The reason for this different behaviour is still not clarified.
142
Figure 11.17.: Heights of DNA structures after different fabrication steps. Topograph
A (7x7 µm) shows all DNA structures after fabrication in STE-buffer. Scheme B helps
to assign the structures to the diagram. Diagram A presents height alterations of the
DNA structures after different fabrication steps.
Remarkable is the dramatic decrease in DNA height when after hybridization (dark
gray bars) a second nanografting step is performed (light gray bars). In this step the
DNA heights are reduced of about 3 to 4.5 nm to a value similar to the situation before
hybridization. In this case a sole solvent induced contraction of the hybridized DNA
molecules is very unlikely. Even if the medium is changed back to STE-buffer (striped
bars) the height decrease is not reversible and only a small increase of about 1-2 nm
is observed. Consequently the height decrease upon a second nanografting step after
hybridization raises the question if the dsDNA denatures during nanografting. In this
context it is interesting to compare the heights of the new ssDNA graftings, produced in
the second nanografting step after hybridization (stripe 2, square 3, square 4), with the
heights of the first ssDNA graftings before hybridization (black and squared bars). It
143
becomes obvious that their heights are very similar. This indicates that only dsDNA is
effected by the reduction in height during the second nanografting step after hybridization. Furthermore the similar heights exclude a change in reference MCH areas. In order
to clarify the effect of the EtOH on the dsDNA additional experiments were carried out
including a compressibility study, SPR experiments and thermal melting measurements.
11.4.2. DNA compressibility study
In general the topography of ”soft” samples, such as SAMs, is measured by scanning
with minimum loading forces to avoid damaging of the SAM and compression of the
molecules. For compressibility studies the relative height is measured as a function of
the applied loading force with respect to a defined reference. Compressibility studies
give information about the mechanical/elastic properties of the probed molecules.
Figure 11.18 presents compressibility measurements of nanografted DNA before and after hybridization and after a second ss-DNA nanografting process. The surrounding
matrix is a MCH SAM. For studying compressibility the loading force was steadily increased after each image scan up to 10.5 nN and finally brought back to the initially low
loading force of 0.3 nN. Height differences in nanostructures are due to different fabrication parameters. Line density for patch 1 is 2.56 and for patch 2 and 3 it is 5.12. Before
hybridization (black curves) DNA shows an approximately linear decrease of height as
response on the applied loading forces. Upon hybridization DNA molecules reorient
which results in an increase in height that was already discussed in section 11.4.1.1. In
addition also the persistence length of DNA extends during hybridization from ∼1 nm
for ssDNA to ∼45 nm for dsDNA. Consequently the molecules become stiffer and their
resistance against an applied force rise (wine curve). Up to a loading force of 2 nN the
height of the DNA remains relatively constant. For loading forces larger than 2 nN a
significant decrease in height occurs.
After a second nanografting of HS-ssDNA subsequently to hybridization again a significant loss in DNA height is detected. But the mechanical response still show a very
similar behaviour than that of dsDNA directly after hybridization (wine curve and red
curve). This indicates that the DNA remains double-stranded and the height decrease
is due to another reason.
It also worth to be noted that the compression of all three nanostructures is fully reversible when reducing the loading force from 10.5 nN back to 0.3 nN.
11.4.3. SPR study on DNA SAMs
Surface plasmon resonance (SPR) spectroscopy is a standard technique for investigating
adsorption and desorption of material onto surfaces. It is sensitive to changes in the
dielectric constant at a metal surface, such as gold. P-polarized monochromatic light is
reflected from a glass prism-metal interface, and the reflectivity of the light is measured
as a function of incident angle (Fig. 11.19). At a certain angle (the plasmon angle), the
evanescent field of the light couples to a plasmon mode that propagates along the metal
144
Figure 11.18.: Compressibility of DNA after different nanofabrication steps. Topographs
present DNA patches surrounded by an MCH SAM in STE buffer at lowest (0.3 nN,
A-C) and highest loading force (10.5 nN, D-F) before and after hybridization and after
a second DNA nanografting process. Graphs 1-3 display compressibility of the DNA
patches in STE buffer.
145
Figure 11.19.: Kretchman-Raether configuration: a beam of p-polarized light is used to
illuminate the backside of a glass slide with a thin film of gold on top covered with a
SAM. At a specific angle ΘSP the evanescent field of the incident light excites plasmons
(collective excitations) of the electrons at the gold surface. At this angle a minimum in
reflected energy is detected. The values of ΘSP change as the index of refraction of the
interface changes (e.g. by the adsorption of molecules). A typical SPR resonance curve
is displayed.
surface, and a minimum in reflected intensity is observed. The plasmon angle (θSP )
is strongly affected by changes in the refractive index at the metal-adsorbate interface
which changes upon adsorption or desorption of target molecules. In a typical SPR
sensorgram the change of the plasmon angle ΘSP over time is used to compute and plot
the change of refractive index as a function of time.
Figure 11.20 A and B (y-axis zoom of graph A) present SPR sensorgrams on a low density DNA SAM. Diagram C summarizes the corresponding changes in index of refraction
after treatments with different reagents. Graph 11.20 A shows large changes of the SPR
signal during injection of EtOH/STE, H2 O and urea. The refractive indices of these
solutions are very different from the refractive index of the STE running buffer. Therefore the SPR signal is temporarily shifted during application of the different solvents. In
literature this effect is also known as ”bulk effect”. The bulk effect has to be separated
from refractive index changes due to adsorption/desorption of molecules at the interface.
In this particular experiments the bulk effect is dominating the adsorption/desorption
processes. Therefore graph B presents an y-axis zoom of graph A focussing on the adsorption/desorption processes. The relative adsorption or desorption of material on the
interface is determined by calculating the difference between the signal before and after
an injection. Graph C summarizes the total amount of adsorbed complementary DNA
at different time points throughout the experiment. Therefore the SPR signal for the
DNA SAM was set to zero in graph C.
First the SAM was hybridized by adding complementary ssDNA (see marker ”1.comp
DNA” in Fig. 11.20 B). Due to the hybridization of surface immobilized and complementary DNA the index of refraction at the interface is changed. Initially a fast adsorption
is observed that slow down at the end of the injection phase. The total amount of hy-
146
Figure 11.20.: SPR data for a low density DNA SAM. A) Index of refraction changes of
a low density DNA SAM upon adsorption of complementary ssDNA and washing with
ethanol/STE-buffer mixture, pure water and 6 M urea. B) Y-axis zoom of graph A. C)
Bar graph of index of refraction changes of a low density DNA SAM after treatment
with different reagents.
147
bridized DNA yields to about 300 µRIU (refractive index units). In order to proof if an
EtOH/STE-buffer mixture affects dsDNA the hybridized DNA SAM was washed with
a 1:1 mixture of EtOH/STE-buffer solution (see marker ”EtOH/STE” in Fig. 11.20 B).
Because the index of refraction of ethanol (n=1.36) is much higher than that of the buffer
(n∼1.342) a large bulk effect is observed (see marker ”EtOH/STE” in Fig. 11.20 A).
After exchange to running buffer the bulk effect is compensated and a decrease in the
SPR signal of about 15% is found which is most likely due to a loss of DNA. When
trying to rehybridize the DNA lost during EtOH/STE washing with a second injection
of complementary DNA the SPR signal exceeds the initial DNA adsorption level (see
marker ”2.comp DNA” in Fig. 11.20 B). This is because the DNA SAM was not saturated with complementary DNA within the first injection of complementary DNA (e.g.
partial hybridization).
Upon washing with pure water a ”negative” bulk effect is observed due to the lower index
of refraction of pure water (n=1.33) in comparison to STE-buffer solution(n∼1.342)(see
marker ”H2 O” in Fig. 11.20 B). After switching back to running buffer the bulk effect
is again compensated and the amount of adsorbed DNA is decreased by about 25%
corresponding to ∼50 µRIU.
In order to demonstrate the reversibility of DNA hybridization the DNA SAM is finally treated with 6 M urea which is known to denature DNA(see marker ”urea” in
Fig. 11.20 B). As expected the signal comes back to its initial value before first application of complementary DNA. A more detailed analysis suggests that the SPR signal falls
even slightly below the initial SPR signal observed for the DNA SAM. This observation
points at a weak desorption of surface tethered DNA.
11.4.4. Melting curves
The most convenient way of monitoring thermal denaturation of DNA is by its ultraviolet (UV) absorbance. When DNA denatures, its UV absorbance increases by about
∼40 % at all wavelengths whereby the shape of the absorbance curve does not change
[33]. Usually changes in DNA absorption upon melting are monitored at 260 nm. The
temperature at which 50% of the DNA strands are in the helical double-stranded state
and the other half are in the ”random-coil” single-stranded state is defined as melting
temperature (Tm ).
Figure 11.21 A depicts the change in absorbance at 260 nm of dsDNA dissolved in a
1:1 ethanol/STE-buffer mixture (black) and a 50% diluted STE-buffer solution (grey).
Dotted curves show the change in absorbance upon heating (melting of duplex DNA)
and solid curves display the cooling down (reassociation of dsDNA). Relative absorbance
is the ratio of the absorbance to that at 20◦ C. Differences among melting and reassociation curves are caused by a delayed temperature control. Temperature is not measured
directly in the solution but at the metallic cuvette holder. Due to thermal resistance the
temperature of the DNA solution and the sample holder are not identic. DNA solution
temperature is always lagging a few degrees behind the sample holder’s temperature.
Therefore during a heating phase the DNA solution is cooler than the temperature
148
11.4.4. Melting curves
Figure 11.21.: A) Melting curves of ds-DNA at 260 nm in a 1:1 mixture of STE-buffer
with ethanol (black) and pure water (grey), respectively. B) UV absorbance spectra of
ds-DNA before (solid) and after thermal treatment (dotted) in both mixtures of STEbuffer at 20◦ C.
recorded at the metallic sample holder while during a cooling phase the situation is vice
versa. When cooling down the resulting DNA is only partially base paired because the
complementary strands will not have sufficient time to find each other since measurement time for one curve is about 60 to 90 min. To this end the cooling down curves do
not reach the initial absorbance value at 20◦ C.
Due to the shift of melting and reassociation curves a good guess for Tm , which is the
temperature at which half of the maximum absorbance is attained, is the average of
Tm for heating and cooling curves. The melting temperature determined in STE-buffer
149
diluted with pure water is 75.5◦ C which is consistent with the calculated one yielding
75.3◦ C. Diluting standard STE- buffer with 1-fold with ethanol Tm dropped to 47.2◦ C.
UV absorbance spectra in Figure 11.21 B are almost equal for all conditions. Variations
before and after melting experiments are most likely due to an incomplete reassociation
of dsDNA (spectra were obtained 30 min after cooling down to 20◦ C) and slight variations in DNA concentration caused by evaporation of small amounts of solvent at high
temperatures.
It can be concluded from the aforementioned data that our 25-mer DNA can acquire
and retain a double-stranded helical structure in a 1:1 mixture of EtOH/STE-buffer,
although its thermal stability is lower than in diluted STE-buffer, as reflected by a
28.3◦ C reduction in the Tm value.
11.4.5. Summarizing discussion on the height decrease of dsDNA
upon nanografting of ssDNA
In subsection 11.4.1.2 a drastic and irreversible decrease in DNA height was observed
when after hybridization (Fig. 11.17, dark gray bars) a second HS-ssDNA nanografting
step was performed (Fig. 11.17, light gray bars). This decrease results in heights similar
to the situation before hybridization. In order to explain this effect a compressibility
study, SPR measurements and UV absorbance spectroscopy were carried out.
Possible causes for the height decrease could be (i) a change in reference height, (ii)
denaturation of duplex DNA, (iii) exchange of immobilized dsDNA with HS-ssDNA
from grafting solution, and (iv) collapse of the dsDNA structure. In the following this
suggestions will be discussed referring to the aforementioned results.
(i) Changes in reference height: A change in MCH reference height is highly unlikely. The height measurements in Figure 11.17 A show that nanografted structures of ssDNA fabricated after hybridization have a very similar height as those
created before hybridization (Fig. 11.17 A, squared vs stripped bars). This would
not be the case if the reference height had changed during sample preparation
steps.
(ii) Denaturation of duplex DNA: Compressibility, SPR and UV measurements indicate that DNA remains at least partly double-stranded despite its decrease in
height. When studying the compressibility of DNA nanopatches after a second
HS-ssDNA nanografting step they still behaves like dsDNA patches (Fig. 11.18).
Up to a force of 1.95 nN dsDNA (wine and red curves) mostly resists a compression by the AFM tip whereas ssDNA (black curves) show less resistance even at
smaller forces of 0.75 nN. Furthermore a destabilization of the duplex DNA caused
by the presence of 50% ethanol in the nanografting solution were disproved by UV
and SPR measurements. Although a concentration of 50% ethanol (v/v) decrease
the melting temperature of dsDNA the duplex is still stable at room temperature
(Fig. 11.21). Nevertheless SPR data indicate denaturation of a small amount of
150
11.4.5. Summarizing discussion on the height decrease of dsDNA upon nanografting of ssDNA
dsDNA molecules (Fig. 11.20). After washing with EtOH/STE solution the SPR
signal drops about approximately 15%.
(iii) Exchange of immobilized dsDNA with HS-ssDNA from grafting solution:
An exchange of surface tethered dsDNA with dissolved HS-ssDNA from solution
might be possible but with regard to the binding energies it is more likely that
some immobilized dsDNA molecules denature, the complementary DNA dissolves
and maybe hybridizes with HS-ssDNA in the grafting solution. This denaturation
could be caused due to the large excess on HS-ssDNA in the nanografting solution
that might shift the hybridization equilibrium between the immobilized dsDNA
and the dissolved HS-ssDNA.
(iv) Collapse of the duplex DNA structure: Recent studies report on more compact configurations of duplex DNA in the presence of ethanol but in general this
effect is reversible when going form ethanolic solution back to buffer solution. The
signal decrease in SPR data after washing with EtOH/STE could also be caused
by a compression of the immobilized dsDNA molecules due to dehydration instead
of a partly denaturation. Maybe the collapsing of duplex DNA is less or slower
reversible than that of ssDNA in ethanolic solution.
Concerning the fact that the height of dsDNA decline to a value very similar to that
of ssDNA it is most likely that the dsDNA areas remain only partly double-stranded.
This is probably due to a simultaneous contribution from different factors discussed
above: denaturation of the immobilized duplex DNA, exchange with molecules from
nanografting solution and collapsing of remaining dsDNA due to dehydration. However,
with respect to the SNIM measurements it could be proofed that DNA remains at least
partly double-stranded.
151
11.5. Characterization of DNA nanostructures using
s-SNIM
The easiest way to test the applicability of s-SNIM in hybridization detection is to fabricate samples with lateral patterns of ssDNA and dsDNA next to each other. Thus both
DNA states can be compared directly under the same conditions (e.g. alignment of the
set-up). For frequency dependent s-SNIM measurements an additional reference field
is necessary which has constant dielectric properties within the investigated frequency
range.
The samples prepared by nanografting provide exactly this requirements. As reference
MCH was selected because it has no prominent absorption features within the investigated spectral region and it is solvable in aqueous buffer and therefore well suited for
nanografting into DNA SAMs. Apart from the detectability of hybridization it is of large
interest to test the ability of s-SNIM to qualify and quantify hybridization efficiencies.
To this end in a first step DNA structures were written into a DNA SAM in order to
compare the near-field signal of DNA SAM and nanografted DNA with each other.
The sample used for this DNA experiments is the same as that characterized by height
measurements in section 11.4.1 (page 139). Its preparation is described in detail in
section 11.2.2.5 (page 121).
Figure 11.22 displays two topographic images (A, C) recorded in dynamic mode in air
simultaneous to s-SNIM measurements. Topograph A shows the nanografted stripes of
DNA and MCH within a DNA SAM and topograph C depicts one of the patch-in-apatch structures (Fig. 11.17 square 2 and 4 (page 143). Diagrams B, D, and E present
corresponding line profiles. In the topographs as well as in the line profiles a clear loss
in height for all structures is observed compared to the contact mode measurements in
STE-buffer (Fig. 11.17, striped bars (page 143)). In air the molecules collapse and stick
together. Nevertheless the trends in height remain the same. For instance the ssDNA
stripe in topograph A is still higher than the dsDNA stripe. The height differences
shown in the line profiles are maximum 1.5 nm (diagram E, height difference between
MCH and ssDNA patch). In the line plots the MCH height was set again to 1.2 nm
with respect to the gold. It has to be considered that most probably also the MCH
collapses in air and that therefore the height referencing with respect to the gold might
not reflect the truth. However, the relative height differences between DNA and MCH
are not affected.
Figure 11.23 presents two schemes (A, E) of the investigated DNA structures. Topographs (B, F) and the corresponding near-field images (C, G) recorded at an IR
frequency of 1679 cm−1 are presented beside the schemes in a linear grey scale. Black
represents minimal height in the topography and minimal detected signal in the nearfield images. All images clearly reveal the nanografted structures. Remarkably is the
near-field contrast of the MCH which is inverse to the topographic contrast. In comparison to the background DNA SAM MCH appears dark in the topographs due to the
152
11.5. Characterization of DNA nanostructures using s-SNIM
Figure 11.22.: Topographic images of patterned DNA and MCH recorded in dynamic
mode in air and depicted in a linear gray scale. B, D, and E show corresponding line
profiles as marked in the topographs. The two schemes help to assign the sample pattern
to the different molecules.
smaller height of MCH. However, it appears bright in the near-field images corresponding to a higher detected infrared signal due to the different dielectric properties of DNA
and MCH molecules. This is most likely because in contrast to the DNA the MCH
molecules show no absorption in the investigated spectral region.
Near-field data were recorded at 7 wavenumbers: ν̃ = 1633, 1644, 1658, 1667, 1679,
1688, and 1722 cm−1 . For calculating the near-field contrast first the signal intensities of
ssDNA, dsDNA, MCH and DNA SAM were determined as an average over each region of
interest (excluding defects in gold). From those the near-field contrast C was calculated
according to the following formula:
C =1−
IDN A
IM CH
(11.1)
For IDN A the corresponding averaged intensities of ssDNA, dsDNA or DNA-SAM were
inserted, respectively. MCH was used as an internal reference because the MCH molecule
153
Figure 11.23.: Schemes (A, E), topographs (B, F) and near-field images (C, G) of two
different investigated DNA patterns. Images (B, C, F, G) are depicted in a linear
gray scale where black represents minimal height and minimal detected infrared signal.
Diagrams D and H present the near-field contrast of each DNA structure at 7 different
wavenumbers. The lines are just guides for the eyes.
has no specific absorption bands within the investigated spectral region (Fig. 11.9). However for the MCH SAM a very small peak was found around 1715 cm−1 in the IRRAS
spectrum in Figure 11.9 which was attributed to physisorbed EDTA. It is expected that
during nanografting the adsorbed EDTA is displaced into solution.
Figure 11.23 D and H display the near-field contrast as a function of the wavenumber.
The lines serve as guides to the eyes and the given errors are the standard deviations
of the averaged intensities. In both diagrams a clear near-field contrast between ssDNA
(solid gray lines) and hybridized DNA (black lines) appear in the spectral region from
1658 to 1688 cm−1 , which is in good agreement with the calculated infrared spectra
(Fig. 11.6, page 129). However even at lower and higher frequencies a very weak contrast is observable for dsDNA as well as for ssDNA. This contrast is likely due to the
fact that even the IRRAS spectra (Fig. 11.10, page 135) and the calculated infrared
spectra (Fig. 11.6, page 129) still show an absorption at these frequencies (excluding the
calculated spectrum for random coil ssDNA). Furthermore topographic artifacts might
contribute to this contrast. As already discussed for the investigated BAT/ODT SAM
in chapter 10 (Fig. 10.8, page 114) also the height differences between MCH and the
DNA can cause an artificial contrast. Due to the strong distance dependence of the
scattered near-field signal with (z+a)3 (see Equation 6.26, page 66) the approximately 1
nm thicker DNA layer causes a reduction of the signal with z/a of ∼2.5%. However, any
artificial contrast based on height differences results in an almost constant offset value
154
11.5. Characterization of DNA nanostructures using s-SNIM
that is independent on wavelength. To this end these contributions can be separated by
measuring the image contrast over several wavelengths.
With regard to an analysis of hybridization efficiency using s-SNIM one has to consider
that the intensity of the near-field signal depends on the number of molecules that are
probed by the evanescent field which evolves from the tip. This number depends on the
one hand on the molecular density and in case of dsDNA on the hybridization efficiency.
Those aspects are illustrated schematically in Figure 11.24. All molecules which contribute to the wavelength dependent near-field contrast are highlighted in red. The more
molecules are highlighted the weaker is the detected infrared signal due to absorption
and the higher is the calculated near-field contrast. In case A and B the hybridization
efficiency is equal but the probe density differs. For B and C the probe density is equal
but the hybridization efficiency is less in case C. The near-field contrast for A and C
is expected to be identical, since for each case one molecule contributes although hybridization efficiency as well as probe density are different. In comparison to A and C
a higher near-field contrast is expected in case B because more molecules are probed by
the evanescent field of the tip.
In general the DNA molecules in nanografted structures are more densely packed than
in a DNA SAM grown from solution. Therefore the almost identical near-field contrast
of nanografted structures (Fig. 11.23 D, H) and hybridized DNA SAM suggest that the
hybridization efficiency is lower in case of nanografted structures. This is very surprising
because Mirmomtaz et al. [79] have recently proven that not as supposed so far the probe
density alone is responsible for hybridization efficiency but mainly the entanglement of
the ssDNA molecules. Since spatially constrained adsorption results in a better ordering
and less entanglement of the ssDNA molecules it is reasonable that the hybridization
efficiency is higher in nanografted structures as in unconstrained assembled monolay-
Figure 11.24.: Dependency of near-field signal intensity on probe density and hybridization efficiency. Molecules contributing to wavelength dependent near-field contrast are
highlighted in red.
155
ers. However, deriving quantitative information about hybridization efficiency from the
s-SNIM results presented above is difficult. The actual loss of dsDNA upon the second
ssDNA nanografting is unknown and might have been different for dsDNA SAM and
nanografted dsDNA.
Taking into account the lateral resolution achieved on the BAT/ODT SAM of (90 nm)2
for the s-SNIM set-up [150] and assuming a molecular density of approximately 1.5 x
1012 immobilized DNA molecules/cm2 [39, 176] and a hybridization efficiency of 50%
[35] yields a lower detection limit of only 61 dsDNA molecules (within an area of (90
nm)2 ).
156
11.6. Summarizing conclusion
11.6. Summarizing conclusion
In this chapter it has been demonstrated that a combination of nanografting with sSNIM is a very promising tool for nanoanalytics.
In the first part different IRRAS experiments on DNA and MCH SAMs were presented
characterizing the infrared spectral response of the DNA sequence employed in this work
when tethered to a gold surface. All experiments were carried out with regard to following s-SNIM measurements.
When comparing SAM spectra of ssDNA and dsDNA IRRAS is a suitable technique for
label free detection of DNA hybridization on gold surfaces. The detected spectra were in
good agreement with the calculated spectra based on the data set of Brewer et al. [166]
although a secondary structure has to be considered for the employed DNA sequence.
IRRAS experiments on the influence of MCH backfilling showed that MCH disrupts nonthiol interactions of the DNA with the gold and therefore improves spectral response.
Since the samples for s-SNIM studies were prepared in the Nanostructure Laboratory at
the ELETTRA in Trieste, Italy also the ageing of DNA SAMs was investigated but no
significant changes in the spectral signature were found.
Using nanografting dsDNA structures, ssDNA structures and MCH reference areas have
been fabricated next to each other. Hybridization of immobilized DNA was detected by
height changes upon formation of duplex DNA. Since a DNA SAM was used as matrix it
was possible to directly compare self-assembled and nanografted DNA monolayers with
each other. Moreover immediately after each fabrication step the AFM was utilized for
characterizing in situ the height changes in the DNA SAM or the nanografted DNA
structures. It was found that dsDNA shows a drastic height decrease when performing
a second ssDNA nanografting subsequently after hybridization. Different experiments
such as compressibility studies, SPR, and UV absorbance have clearly shown the stability of duplex DNA in a 1:1 mixture of ethanol and STE-buffer solution that was used
for HS-ssDNA nanografting. Thus, the height decrease of immobilized dsDNA upon
nanografting of ssDNA can only be due to an exchange with molecules from nanografting
solution, a collapsing of remaining dsDNA due to dehydration or due to a shifted duplex
equilibrium caused by the excess of HS-ssDNA in the nanografting solution. However
most likely an interplay of all factors is responsible for the observed effect. Determination of the most crucial factor causing the height decrease needs more experiments and
goes beyond the scope of this thesis. With respect to the s-SNIM measurements it was
important to prove that the DNA remains at least partly in its double-stranded state
after a second HS-ssDNA nanografting which has been demonstrated successfully using
a couple of different techniques.
It is demonstrated that s-SNIM is able to distinguish between single-stranded and
double-stranded DNA in SAMs and also in nanostructures. Therefore s-SNIM is a
promising tool for a qualitative analysis of hybridization at the nanoscale. With regard to quantification of hybridization efficiency further studies on DNA with different
packing densities are necessary. To this end especially nanografting is a well suited
structuring technique because the probe density can be systematically varied during the
writing process [79].
157
12. Infrared dyes in s-SNIM
Contents
12.1. Introduction
. . . . . . . . . . . . . . . . . . . . . . . . . . . . 160
12.2.1. Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
160
12.2.2. Preparation of homogeneous SAMs for IRRAS measurements
160
12.2.3. Preparation of microstructured SAMs . . . . . . . . . . . . .
160
161
12.2.5. s-SNIM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
161
12.3. Results and Discussion . . . . . . . . . . . . . . . . . . . . . . 161
12.4. Conclusion
. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 165
159
Chapter 12. Infrared dyes in s-SNIM
12.1. Introduction
Metal carbonyl compounds [My (CO)x ] have specific and intense CO stretching vibrations
within 2150-1900 cm−1 . Therefore they are ideal markers for infrared spectroscopy
because natural organic molecules do not have vibrational features in this region.
12.2.1. Reagents
The cymantrene labeled peptide (CymPntCys) was synthesized from Dr. Harmel Peindy
in the group of Prof. Dr. N. Metzler-Nolte (Ruhr-University of Bochum). A molecular
structure of the molecule is displayed in Figure 12.1. 1-octadecanethiol (ODT, Aldrich,
98%) was purchased from Sigma. Ethanol was used in p.a. quality. Aqueous solutions
were prepared using HPLC water (J.T.Baker).
Figure 12.1.: Bond-line structure of CymPntCys. In red the infrared active cymantrene
group and in blue the thiol group.
12.2.2. Preparation of homogeneous SAMs for IRRAS
measurements
Sputter deposited gold on silicon (Anfatec Instruments AG) was used as substrate (see
chapter 7 (page 75)). Prior to use substrates were cleaned with hot piranha solution (7:3
mixture of 96% H2 SO4 and 30% H2 O2 ) for approximately 15 min (detailed description
see section 7.1.1 (page 76)). Homogeneous SAMs were prepared by immersing clean
gold substrates into 1 mM ethanolic or aqueous solution of CymPntCys. Afterwards the
substrates were rinsed with the respective solvent and dried in a stream of nitrogen.
Laterally structured SAMs of ODT and CymPntCys were fabricated using microcontact
printing. Microcontact printing were performed as described in chapter 8 (page 83).
160
Briefly, prior to use a stamp, fabricated from a TGG01 calibration grid (Mikromasch),
was cleaned and subsequently loaded with ODT by immersion in a 5 mM ethanolic
solution of ODT. Printing was done by gently pressing the dried stamp on a clean TSG
substrate. After a contact time of 120 s the stamp was carefully peeled off. Immediately
after printing the substrate was immersed into a 1 mM ethanolic solution of CymPntCys
over night (> 16 h) in order to fill the remaining bare gold areas with cymantrene
peptide. Afterwards the substrate was thoroughly rinsed with absolute ethanol and
dried in a stream of nitrogen.
IRRAS spectra were recorded on a Vertex V80 (Bruker) spectrometer equipped with a
globar source, a KBr beam splitter, a BaF2 polarizer (LOT), a grazing incident reflection
unit and a liquid nitrogen cooled MCT detector. The gold grid polarizer was used to
obtain p-polarized radiation before reflecting off the sample at an incident angle of 80◦
from the surface normal. Spectra were recorded at room temperature with a resolution
of 4 cm−1 and a coadding of 256 spectra. Fourier transformation of all spectra was
carried out with Mertz phase correction and triangular apodization. Evacuation time
for the sample chamber was 5 min. As reference a bare gold substrate was used. Spectra
were evaluated using OPUS-Software 6.5 (Bruker).
12.2.5. s-SNIM
s-SNIM measurements were carried out as described in chapter 9 (page 91). Each image
was recorded at a scan rate of 0.5 Hz and a time constant of 1 ms at the lock-in amplifier.
Image Processing
Image processing was performed using WSxM imaging software 4.0 Develop 10.4 (Nanotec Electronica) [167]. Raw images were corrected using a linear flatten function and
for presentation the contrast was enhanced.
Figure 12.2 presents IRRAS spectra of CymPntCys on gold adsorbed from ethanolic
solution (panel A) or aqueous solution (panel B). For both solutions three different immobilization times 1 h, 17 h and 23 h were investigated.
All spectra show characteristic vibrational bands for the cymantrene group
(C5 H2 −Mn−C≡O ↔ C5 H2 −Mn=C=O) at ∼2028 cm−1 and ∼1949 cm−1 . Furthermore the amide I band around 1660 cm−1 and the amide II band around 1515 cm−1 are
clearly visible. Except for the 1 h spectrum in panel A the C-H vibration bands around
3000 cm−1 are very weak. This high absorption band in the upper spectrum of panel A
appear only transiently and are probably vanishing due to reorientation of the molecules
during SAM formation.
161
Figure 12.2.: IRRAS spectra of CymPntCys adsorbed on gold from ethanolic solution
(A) or aqueous solution (B). Immobilization times were 1, 17, and 23 hours. Spectra
are not background corrected.
162
When comparing in panel A 1 h and 17 h of incubation the ratio between amide and
metal carbonyl absorption bands is changed. With longer incubation times the amide
peak becomes more pronounced e.g. at 17 h the amide peak is 3 times higher than the
metal carbonyl vibration peak. However, when adsorbed from aqueous solution (panel
B) the amide I peak does not increase over time. The intensities of the amide I and
amide II peak remain very similar.
When comparing the immobilization from ethanolic solution with that from aqueous
solution the metal carbonyl vibrations are more intense (∼1.5 times) in the latter case.
In addition it should be noticed that in both cases incubation times of 1 hour is not
sufficient enough and it is advisable to grow the peptide SAMs over night.
In chapter 11.3.2.1 (page 131) the solvent was already discussed as crucial factor for
SAM formation of MCH molecules. The growth of peptide SAMs seems to be another
example where the SAM formation is affected by the solvent. However the modulation
of the peptide SAM spectra is not as pronounced as with MCH-SAMs. Based on the
spectroscopical findings it was decided absorbing the peptide molecules from ethanolic
solution during the experiments with laterally structured ODT SAMs.
Prior to the experiments with laterally structured SAMs it was investigated if peptide
molecules do unspecifically adsorb to ODT SAM. To this end an ODT SAM was grown
for 4 h and then incubated over night in a solution containing 1 mM of the peptide.
figure 12.3 displays no changes for the ODT spectrum before and after incubation in the
Figure 12.3.: Investigation of unspecific adsorption of CymPntCys on an ODT SAM
163
Figure 12.4.: A) AFM topography image of a laterally structured ODT-CymPntCys
SAM. B) Corresponding height profile averaged over 7 lines to increase signal-to-noise
ratio.
peptide solution. This indicates that within the sensitivity of the FTIR spectrometer no
peptide molecules are unspecifically adsorbed to the ODT SAM. Please note that there
is also no exchange of thiols over 16 h of incubation.
In Figure 12.4 a topographic AFM image of a laterally structured SAM composed of
ODT and CymPntCys molecules is presented. Stripes appearing bright are assigned to
ODT covered regions while the darker areas belong to peptide covered regions. The corresponding height profile shows periodic changes in the height of the adsorbed molecules.
For well ordered ODT molecules stamped onto clean gold a step height of about 2.5 nm
is expected. Since the topography profile gives still a height of 2 nm the peptide SAM
cannot be densely packed. Most likely the density of the peptide molecules is so low
that the molecules are lying down onto the gold. Maybe the side chain groups (e.g.
Lys, Tyr) are promoting intensive interactions with the gold surface. The bright dots
statistically distributed over the image could be either aggregates of peptide molecules
or contamination from solution.
The laterally structured ODT-peptide SAM surface were found ideally suited for characterization with s-SNIM measurements. As Figure 12.5 depicts the laser source was
tuned to three distinct frequency regimes. First 3 different wavenumbers close to the
absorption peak of the metal carbonyl species were investigated. Then a non absorbing wavenumber located between metal carbonyl and amide absorption peak was chosen.
And finally the center wavenumber of the amide absorption peak was studied by s-SNIM.
All s-SNIM images that were recorded at the center of the absorption peaks clearly reveal
the periodic pattern. The observed periodicity is consistent with the printed pattern.
The image recorded at 1931 cm−1 show a very weak contrast. Only the image far away
from the absorption peaks displays no periodic contrast.
164
12.4. Conclusion
Figure 12.5.: IRRAS spectrum of CymPntCys on gold with corresponding near-field
images at five different wavenumbers (colored images). A typical topograph (gray) is
displayed at the top.
12.4. Conclusion
Although the chosen cymantrene labeled peptide is not ideal for growing densely packed
and well structured SAMs the presented measurements are very promising. Despite
of the weak ordering it was still possible to resolve periodic contrasts as a function of
the wavenumber in the s-SNIM measurements. In the IRRAS experiments the strong
absorption of the metal carbonyl group is recognized. Future studies should aim at
improving the ordering and coupling of the peptide towards the surface. It is quite
likely that this will result in even stronger absorption signals. From the presented set of
experiments it cannot be excluded that the metal carbonyl group is in close proximity
of the gold. In combination with DNA molecules the labeling can help to quantify
hybridization efficiencies. A weak point of this methodology is that the initial benefit of
s-SNIM as being a label-free method is lost.
165
13. Bibliography
[1] J. Christopher Love, Lara A. Estroff, Jennah K. Kriebel, Ralph G. Nuzzo, and
Geogre M. Whitesides. Chemical Reviews, 105:1103–1169, 2005.
[2] I. Langmuir. J. Am. Chem. Soc., 39:1848–1906, 1917.
[3] K. Blodgett. J. Am. Chem. Soc., 57:1007–1022, 1935.
[4] W. C. Bigelow, D. L. Pickett, and W. A. Zisman. J. Colloid Interface Sci, 1:513,
1946.
[5] J. Sagiv. J. Am. Chem. Soc., 102(1):92–98, 1980.
[6] D. L. Allara and R. G. Nuzzo. Langmuir, 1:45–52, 1985.
[7] R. G. Nuzzo and D. L. Allara. J. Am. Chem. Soc., 105:4481–4483, 1983.
[8] F. Schreiber. J. Phys.: Condens. Matter, 16:R881–R900, 2004.
[9] R. G. Nuzzo, F. A. Fusco, and D. L. Allara. J. Am. Chem. Soc., 109(8):2358–2368,
1987.
[10] K. Masahiro and S. Noboru. J. Mater. Sci., 28:5088–5091, 1993.
[11] Qian Tang, San-Qiang Shi, and Li-min Zhou.
5(12):2167–2171, 2005.
J. Nanosci. Nanotechnol.,
[12] Stephen E. Creager, Lisa A. Hockett, and Gary K. Rowe. Langmuir, 8:854–861,
1992.
[13] Ajit K. Mahapatro, Adina Scott, Allene Manning, and David B. Janes. Applied
Physics Letters, 88:151917, 2006.
[14] Martin Hegner, Peter Wagner, and Giorgio Semenza. Surface Science, 291:39–46,
1993.
[15] Dimitris Stamou, Delphine Gourdon, Martha Liley, Nancy A. Brunham, Andrzej
Kulik, Horst Vogel, and Claus Duschl. Langmuir, 13:2425–2428, 1997.
[16] J. Diebel, H. Löwe, P. Samori, and J.P. Rabe. Applied Physics A: Materials Science
and Processing, 73(3):273–279, 2001.
167
Bibliography
[17] Peter Wagner, Martin Hegner, Hans-Joachim Güntherodt, and Giorgio Semenza.
Langmuir, 11:3867–3875, 1995.
[18] David W. Mosley, Brian Y. Chow, and Joseph M. Jacobson. Langmuir, 22:2437–
2440, 2006.
[19] C. D. Bain, B. Troughton, Y.-T. Tao, J. Evall, G. M. Whitesides, and R. G. Nuzzo.
J. Am. Chem. Soc., 111:321–335, 1989.
[20] T. Smith. Journal of Colloid and Interface Science, 75(1):51–55, 1980.
[21] M. Schneegans and E. Menzel. Journal of Colloid and Interface Science, 88(1):97–
99, 1982.
[22] A. Ulman. Chem. Rev., 96:1533–1554, 1996.
[23] L. Strong and G. M. Whitesides. Langmuir, 4:546–558, 1988.
[24] C. A. Widrig, C. A. Alves, and M. D. Porter. J. Am. Chem. Soc., 113(8):2805–
2810, 1991.
[25] C. A. Widrig, C. Chung, and M. D. Porter. J. Electroanalytical Chemistry, 310(12):335–359, 1991.
[26] C. E. D. Chidsey and D. N. Loiacono. Langmuir, 6(3):682–691, 1990.
[27] E. Ostuni, L. Yan, and G.M. Whitesides. Colloids and Surfaces, B: Biointerfaces,
15(1):3–30, 1999.
[28] L. H. Dubois and R. G. Nuzzo. Annu. Rev. Phys. Chem., 43:437–463, 1992.
[29] Marc D. Porter, Thomas B. Bright, David L. Allara, and Christopher E. D. Chidsey. J. Am. Chem. Soc., 109:3559–3568, 1987.
[30] J. Spinke, M. Liley, F.-J. Schmitt, H.-J. Guder, L. Angermaier, and B. Knoll. J.
Chem. Phys., 99(9):7012–7019, 1993.
[31] J. L. Wilbur, A. Kumar, H. A. Biebuyck, K. Enoch, and G. M. Whitesides. Nanotechnology, 7:452–457, 1996.
[32] Simon Flink, Frank C. J. M. van Veggel, and David N. Reinhoudt. Adv. Mater.,
12(18):1315–1328, 2000.
[33] D. Voet and J. G. Voet. Biochemistry. John Wiley and Sons, Inc., 1995.
[34] C. Chen, W. Wang, Z. Wang, F. Wei, and X. S. Zhao. Nucleic Acids Res.,
35(9):2875–2884, 2007.
[35] Yang Gao, Lauren K. Wolf, and Rosina M. Georgiadis. Nucleic Acids Research,
34(11):3370–3377, 2006.
168
Bibliography
[36] M. Zuker. Nucleic Acids Res., 31(13):3406–3415, 2003.
[37] D. V. Leff, L. Brandt, and J. R. Heath. Langmuir, 12:4723–4730, 1996.
[38] C. Xu, L. Sun, L. J. Kepley, and R. M. Crooks. Anal. Chem., 65:2102–2107, 1994.
[39] T. M. Herne and M. J. Tarlov. J. Am. Chem. Soc., 119:8916–8920, 1997.
[40] A. B. Steel, R. L. Levicky, T. M. Herne, and M. J. Tarlov. Biophys. J., 79:975–981,
2000.
[41] S. D. Keighley, P. Li, P. Estrela, and P. Migliorato. Biosensors and Bioelectronics,
23:1291–1297, 2008.
[42] Alexander W. Peterson, Richard J. Heaton, and Rosina M. Georgiadis. Nucleic
Acids Research, 29(24):5163–5168, 2001.
[43] D.Y. Petrovykh, H. Kimura-Suda, L.J. Whitman, and M.J. Tarlov. J. Am. Chem.
Soc., 125:5219–5226, 2003.
[44] J. F. Marko and S. Cocco. Physics World, March:37–41, 2003.
[45] M. Liu, N. A. Amro, C. S. Chow, and G.-y. Liu. Nano Letters, 2(8):863–867, 2002.
[46] J. Bednar, P. Furrer, V Katritch, A. Z. Stasiak, J. Dubochet, and A. J. Stasiak.
J. Mol. Biol., 254:579–594, 1995.
[47] Peter Haring Bolivar, Michael Nagel, Frank Richter, Martin Brucherseifer, Heinrich Kurz, Anja Bosserhoff, and Reinhard Büttner. Phil. Trans. R. Soc. Lond. A,
362:323–335, 2004.
[48] A. Kumar and G.M. Whitesides. Appl. Phys. Lett., 63:2002–2004, 1993.
[49] Y. Xia and G. M. Whitesides. Annu. Rev. Mater. Sci., 28:153–184, 1998.
[50] G. M. Whitesides, E. Ostuni, S. Takayama, X. Jiang, and D. E. Ingber. Annu.
Rev. Biomed. Eng., 3:335–373, 2001.
[51] E. Delamarche, H. Schmid, B. Michel, and H. Biebuyck. Adv. Mater., 9:741–746,
1997.
[52] H. Schmid and B. Michel. Macromolecules, 33(8):3042–3049, 2000.
[53] G. Csucs, T. Künzler, K. Feldman, F. Robin, and N. D. Spencer. Langmuir,
19:6104–6109, 2003.
[54] C. D. James, R. C. Davis, L. Kam, H. G. Craighead, M. Isaacson, J. N. Turner,
and W. Shain. Langmuir, 14:741–744, 1998.
169
Bibliography
[55] T. W. Odom, J. Ch. Love, D. B. Wolfe, K. E. Paul, and G. M. Whitesides. Langmuir, 18:5314–5320, 2002.
[56] R. R. Fuierer, R. L. Carroll, D. L. Feldheim, and C. B. Gorman. Adv. Mater.,
14(2):154–157, 2002.
[57] R. D. Piner, J. Zhu, F. Xu, S. Hong, and C. A. Mirkin. Science, 283:661–663,
1999.
[58] M. Peter, X.-M. Li, J. Huskens, and D. N. Reinhoudt. J. Am. Chem. Soc.,
126:11684–11690, 2004.
[59] S. Krämer, R. R. Fuierer, and C. B. Gorman. Chem. Rev., 103:4367–4418, 2003.
[60] A. A. Tseng, A. Notargiacomo, and T. P. Chen. J. Vac. Sci. Technol. B, 23(3):877–
894, 2005.
[61] R. Garcia, R. V. Martinez, and J. Martinez. Chem. Soc. Rev., 35:29–38, 2006.
[62] S. Xu and Gang-yu Liu. Langmuir, 13:127–129, 1997.
[63] M. Liu, N. A. Amro, and G.-y. Liu. Annu. Rev. Phys. Chem., 59:367–386, 2008.
[64] G.-y. Liu, S. Xu, and Y. Qian. Acc. Chem. Res., 33:457–466, 2000.
[65] S. Xu, P. E. Laibinis, and G.-y. Liu. J. Am. Chem. Soc., 120(36):9356–9361, 1998.
[66] S. Xu, S. Miller, P. E. Laibinis, and G.-y. Liu. Langmuir, 15:7244–7251, 1999.
[67] J.-j. Yu. Agilent Technologies - Application Note, 5989-7699EN, 2007.
[68] G.-y. Liu and N. A. Amro. PNAS, 99(8):5165–5170, 2002.
[69] K. Salaita, Y. Wang, J. Fragala, R. A. Vega, C. Liu, and C. A. Mirkin. Angew.
Chem. Int. Ed., 45:7220–7223, 2006.
[70] M. A. Case, G. L. McLendon, Y. Hu, T. K. Vanderlick, and G. Scoles. Nano
Letters, 3(4):425–429, 2003.
[71] Y. Hu, A. Das, M. H. Hecht, and G. Scoles. Langmuir, 21(20):9103–9109, 2005.
[72] Ch. Staii, D. W. Wood, and G. Scoles. J. Am. Chem. Soc., 130(2):640–646, 2008.
[73] Ch. Staii, D. W. Wood, and G. Scoles. Nano Letters, 8(8):2503–2509, 2008.
[74] K. Wadu-Mesthrige, S. Xu, N. A. Amro, and G.-y. Liu. Langmuir, 15:8580–8583,
1999.
[75] K. Wadu-Mesthrige, N. A. Amro, J. C. Garno, S. Xu, and G.-y. Liu. Biophys. J.,
80:1891–1899, 2001.
170
Bibliography
[76] D. Zhou, X. Wang, L. Birch, R. Trevor, and C. Abell. Langmuir, 19(25):10557–
10562, 2003.
[77] J. Ngunjiri and J. C. Garno. Analytical Chemistry, 80(5):1361–1369, 2008.
[78] M. Liu and G.-y. Liu. Langmuir, 21:1972–1978, 2005.
[79] E. Mirmomtaz, M. Castronovo, Ch. Grunwald, F. Bano, D. Scaini, A. A. Ensafi,
G. Scoles, and L. Casalis. Nano Lett., 2008.
[80] C. D. Frisbie, L. F. Rosznyai, A. Noy, M. S. Wrighton, and C. M. Lieber. Science,
265:2071, 1994.
[81] A. Noy, D. V. Vezenov, and C. M. Lieber. Annu. Rev. Mater. Sci., 27:381–421,
1997.
[82] Inc. Hitachi High Technologies America.
[83] Y. Garini, B. J. Vermolen, and I. T. Young. Current Opinion in Biotechnology,
16:3–12, 2005.
[84] S. W. Hell and E. H. K. Stelzer. Opt. Commun., 93:277–282, 1992.
[85] S. W. Hell and J. Wichmann. Opt. Lett., 19:780–782, 1994.
[86] E.H. Synge. Philosophical Magazine (1798-1977), 6:356–62, 1928.
[87] H. A. Bethe. Phys. Rev., 66:163–182, 1944.
[88] C. Bouwkamp. Philips Res. Report, 5:321–332, 1950.
[89] C. Bouwkamp. Philips Res. Rep., 5:401–422, 1950.
[90] J. A. O’Keefe. J. Opt. Soc. America, 46(5):359, 1956.
[91] C. W. McCutchen. J. Am. Chem. Soc., 57:1190–1192, 1967.
[92] E. A. Ash and G. Nicholls. Nature, 237:510–512, 1972.
[93] G. Binning, H. Rohrer, Ch. Gerber, and E. Weibel. Phys. Rev. Lett., 49(1):57–61,
1982.
[94] D. W. Pohl, W. Denk, and M. Lanz. Appl. Phys. Lett., 44(7):651–653, 1984.
[95] A. Lewis, M. Isaacson, A. Harootunian, and A. Muray. Ultramicroscopy, 13:227–
232, 1984.
[96] E. Betzig, J. K. Trautman, T. D. Harris, J. S. Weiner, and R. L. Kostelar. Science,
251:1468–1470, 1991.
[97] J. Wessel. J. Opt. Soc. America B, 2(9):1538–1541, 1985.
171
Bibliography
[98] A. Lahrech, R. Bachelot, P. Gleyzes, and A. C. Boccara.
21(17):1315–1317, 1996.
Optics Letters,
[99] B. Knoll and F. Keilmann. Appl. Phys. A, 66:477–481, 1998.
[100] N. C. J. v.d. Valk and P. C. M. Planken. Appl. Phys. Lett., 81(9):1558–1560, 2002.
[101] F. Keilmann, D. W. v.d. Weide, T. Eickelkamp, R. Merz, and D. Stöckle. Opt.
Commun., 129:15–18, 1996.
[102] L. Novotny, D. W. Pohl, and B. Hecht. Optics Letters, 20(9):970, 1995.
[103] M. K. Hong, A. G. Jeung, N. V. Dokholyan, T. I. Smith, H. A. Schwettman,
P. Huie, and S. Erramilli. Nuclear Instruments and Methods in Physics Research
B, 144:246–255, 1998.
[104] B. Hecht, B. Sick, U. P. Wild, V. Deckert, R. Zenobi, O. J. F. Martin, and D. W.
Pohl. J. Chem. Phys., 112(18):7761–7774, 2000.
[105] A. Cricenti, R. Generosi, P. Perfetti, J. M. Gilligan, N. H. Tolk, C. Coluzza, and
G. Margaritondo. Appl. Phys. Lett., 73(2):151–153, 1998.
[106] A. Cricenti, R. Generosi, M. Barchesi, M. Luce, and M. Rinaldi. Rev. Sci. Instrum.,
69(9):3240–3244, 1998.
[107] A. Cricenti, R Generosi, M. Luce, P. Perfetti, G. Margaritondo, D. Talley, J. S.
Sanghera, I. D. Aggarwal, N. H. Tolk, A. Congiu Castellano, M. A. Rizzo, and
D. W. Piston. Biophys. J., 85:2705–2710, 2003.
[108] P. Ephrat, K. Roodenko, L. Nagli, and A. Katzir. Appl. Phys. Lett., 84:637, 2004.
[109] M. Platkov, A. Tsun, L. Nagli, and A. Katzir. Rev. Sci. Instrum., 77:126103, 2006.
[110] F. Zenhausern, Y. Martin, and H. K. Wickramasinghe. Science, 269(5227):1083–
1085, 1995.
[111] D. Haeflinger, J. M. Plitzko, and R. Hillenbrand. Appl. Phys. Lett., 85(19):4466–
4468, 2004.
[112] T. Yamaguchi, S. Yoshida, and A. Kinbara. Thin Solid Films, 21:173–187, 1974.
[113] B. Knoll and F. Keilmann. Opt. Commun., 182:321–328, 2000.
[114] S. G. Moiseev and S. V. Sukhov. Optics and Spectroscopy, 98(2):308–313, 2005.
[115] J. Aizpurua, T. Taubner, F. J. G. de Abajo, M. Brehm, and R. Hillenbrand. Optics
Express, 16(3):1529–1545, 2008.
[116] R. Hillenbrand, B. Knoll, and F. Keilmann. Journal of Microscopy, 202(1):77–83,
2001.
172
Bibliography
[117] R. Hillenbrand and F. Keilmann. Phys. Rev. Lett., 85(14):3029–3032, 2000.
[118] E. Bründermann and M. Havenith. Annu. Rep. Prog. Chem., Sect. C Phys. Chem.,
104:235–255, 2008.
[119] L. Gomez, R. Bachelot, A. Bouhelier, G. P. Wiederrecht, S. Chang, S. K. Gray,
F. Hua, S. Jeon, J. A. Rogers, M. E. Castro, S. Blaize, I. Stefanon, G. Lerondel,
and P. Royer. J. Opt. Soc. Am. B, 23:823–833, 2006.
[120] T. Taubner, R. Hillenbrand, and F. Keilmann. Journal of Microscopy, 210:311–
314, 2003.
[121] F. de Lange, A. Cambi, R. Huijbens, B. de Bakker, W. Rensen, M. Garcia-Parajo,
N. van Hulst, and C. G. Figdor. Journal of Cell Science, 114(23):4153–4160, 2001.
[122] M. Koopman, A. Cambi, B. I. de Bakker, B. Joosten, C. G. Figdor, N. F. Hulst,
and M. F. Garcia-Parajo. FEBS Letters, 573:6–10, 2004.
[123] C. Höppner, D. Molenda, H. Fuchs, and A. Naber. Journal of Microscopy, 210:288–
293, 2003.
[124] J. A. De Aro, K. D. Weston, S. K. Burrato, and U. Lemmer. Chem. Phys. Lett.,
277:532–538, 1997.
[125] A. Cadby, R. Dean, A. M. Fox, R. A. Jones, and D. G. Lidzey. Nano Lett.,
5:2232–2237, 2005.
[126] R. Stevenson, R. Riehn, and R. G. Milner. Appl. Phys. Lett., 79:833–835, 2001.
[127] R. Pomraenke, C. Ropers, J. Renard, C. Lienau, L. Lüer, D. Polli, and G. Cerullo.
Journal of Microscopy, 229(2):197–202, 2008.
[128] A. Hartschuh, E. J. Sanchez, X. S. Xie, and L. Novotny. Phys. Rev. Lett., 90:95503,
2003.
[129] N. Anderson, P. Anger, A. Hartschuh, and L. Novotny. Nano Lett., 6(4):744–749,
2006.
[130] U. Neugebauer, P. Rösch, M. Schmitt, J. Popp, C. Julien, A. Rasmussen, C. Budich, and V. Deckert. Chem Phys Chem, 7(7):1428–1430, 2006.
[131] U. Neugebauer, U. Schmid, K. Baumann, W. Ziebuhr, S. Kozitskaya, V. Deckert,
M. Schmitt, and J. Popp. Chem Phys Chem, 8(1):124–137, 2007.
[132] B.-S. Yeo, S. Mädler, T. Schmid, W. Zhang, and R. Zenobi. J. Phys. Chem. C,
112:4867–4873, 2008.
[133] E. Bailo and V. Deckert. Angew. Chem. Int. Ed., 47(9):1658–1661, 2008.
173
Bibliography
[134] M. Platkov, A. Tsun, L. Nagli, and A. Katzir. Appl. Phys. Lett., 92:104104, 2008.
[135] J. Generosi, G. Margaritondo, J. S. Sanghera, I. D. Aggarwal, N. H. Tolk, D. W.
Piston, A. Congiu Castellano, and A. Cricenti. Journal of Microscopy, 229(2):259–
263, 2008.
[136] B. Dragnea, J. Preusser, J.M. Szarko, L. A. McDonough, S.R. Leone, and W.D.
Hinsberg. Applied Surface Science, 175-176:783–789, 2001.
[137] C. A. Michaels, X. Gu, D. B. Chase, and S. J. Stranick. Appl. Spectroscopy,
58(3):257–263, 2004.
[138] L. A. McDonough, B. Dragnea, J. Preusser, S.R. Leone, and W.D. Hinsberg.
Journal of Physical Chemistry B, 107(21):4951–54, 2003.
[139] T. Taubner, F. Keilmann, and R. Hillenbrand. Optics Express, 13(22):8893–8899,
2005.
[140] A. Lahrech, R. Bachelot, P. Gleyzes, and A. C. Boccara. Appl. Phys. Lett.,
71(5):575–577, 1997.
[141] B. Knoll and F. Keilmann. Appl. Phys. Lett., 77(24):3980–3982, 2000.
[142] J.-S. Samson, G. Wollny, E. Bründermann, A. Bergner, A. Hecker, G. Schwaab,
A.D. Wieck, and M. Havenith. Phys. Chem. Chem. Phys., 8(6):753–58, 2006.
[143] B.B. Akhremitchev, S. Pollack, and G.C. Walker. Langmuir, 17:2774–2781, 2001.
[144] M. B. Raschke, L. Molina, T. Elsaesser, D. H. Kim, W. Knoll, and K. Hinrichs.
Chem Phys Chem, 6:2197–2203, 2005.
[145] T. Taubner, R. Hillenbrand, and F. Keilmann. Appl. Phys. Lett., 85(21):5064–66,
2004.
[146] M. Brehm, T. Taubner, R. Hillenbrand, and F. Keilmann. Nano Letters, 6(7):1307–
1310, 2006.
[147] G. Wollny, E. Bründermann, Z. Arsov, L. Quaroni, and M. Havenith. Optics
Express, 16(10):7453–7459, 2008.
[148] G. Wollny. Infrarot-Nahfeldmikroskopie von Phospholipiden. PhD thesis, RuhrUniversität Bochum, 2008.
[149] B.B. Akhremitchev, Y. Sun, L. Stebounova, and G.C. Walker.
18(14):5325–5328, 2002.
Langmuir,
[150] I. Kopf, J.-S. Samson, G. Wollny, Ch. Grunwald, E. Bründermann, and
M. Havenith. J. Phys. Chem. C, 111:8166–8171, 2007.
174
Bibliography
[151] G. C. Cho, H.-T. Chen, S. Kraatz, N. Karpowicz, and R. Kersting. Semicond. Sci.
Technol., 20:S286–S292, 2005.
[152] A. J. Huber, F. Keilmann, J. Wittborn, J. Aizpurua, and R. Hillenbrand. Nano
Lett., 2008.
[153] Ralf Arnold. Struktur und Ordnung selbstordnender Monolagen aliphatischer und
aromatischer Thiole auf Goldoberflächen. PhD thesis, Ruhr-Universität, 2001.
[154] Andreas Bergner. Infrarot Mikro- und Nanomikrospektroskopie. PhD thesis, RuhrUniversität Bochum, 2004.
[155] D. W. Lynch and W. R. Hunter. Handbook of optical constants of solids. 1985.
[156] Andreas Hecker.
Entwicklung optisch parametrischer Oszillatoren zur
hochauflösenden Spektroskopie und Analyse des van-der-Waals-Komplexes
(N2O)2. PhD thesis, Ruhr-Universität Bochum, 2003.
[157] C. K. N. Patel. Appl. Phys. Lett., 7:246–247, 1965.
[158] C. E. Treanor, J. W. Rich, and R. G. Rehm. J. Chem. Phys., 48(4):1798–1807,
1968.
[159] C. Freed. Appl. Phys. Lett., 18:458–461, 1971.
[160] M. Gromoll-Bohle, W. Bohle, and W. Urban. Opt. Commun., 69(5-6):409–413,
1989.
[161] E. Bachem, A. Dax, T. Fink, A. Weidenfeller, M. Schneider, and W. Urban. Appl.
Phys. B: Lasers Opt., 57(3):185–191, 1993.
[162] W. Urban. Infrared Phys. Technol., 36(1):465–473, 1995.
[163] Angelika Schubert. Untersuchung am sealed-off CO-Laser mit variablem Temperaturübergang. PhD thesis, Universität Bonn, 1994.
[164] M. Hesse, H. Meier, and B. Zeeh. Spektroskopische Methoden in der organischen
Chemie. George Thieme Verlag, 1995.
[165] Christian Grunwald. Proteinadsorption an organischen Modelloberflächen. PhD
thesis, Ruhr-Universität, 2005.
[166] Scott H. Brewer, Selina J. Anthireya, Simon E. Lappi, David L. Drapcho, and
Stefan Franzen. Langmuir, 18:4460–4464, 2002.
[167] I. Horcas, R. Fernandez, Gomez-Rodriguez, J. Colchero, J. Gomez-Herrero, and
A. M. Baro. Rev. Sci. Instrum., 78(013705):1–8, 2007.
[168] K.-I. Miyamoto, K. Onodera, R. Yamaguchi, K. Ishibashi, Y. Kimura, and M. Niwano. Chem. Phys. Lett., 436:233–238, 2007.
175
Bibliography
[169] Ralph G. Nuzzo, Lawrence H. Dubois, and David L. Allara. J. Am. Chem. Soc.,
112:558–569, 1990.
[170] K. A. Peterlinz and R. M. Georgiadis. Langmuir, 12:4731–4740, 1996.
[171] O. Dannenberger, J. J. Wolff, and M. Buck. Langmuir, 14:4679–4682, 1998.
[172] D. Yan, J. A. Saunders, and G. K. Jennings. Langmuir, 19:9290–9296, 2003.
[173] K. Tsukamoto, T. Kubo, and H. Nozoye. Applied Surface Science, 244:578–583,
2005.
[174] R. Yamaguchi, K. Miyamoto, K. Ishibashi, and A. Hirano.
102:014303, 2007.
J. Appl. Phys.,
[175] R. Levicky, T. M. Herne, M. J. Tarlov, and S. K. Satija. J. Am. Chem. Soc.,
120:9787–9792, 1998.
[176] A. W. Peterson, Lauren K. Wolf, and R. M. Georgiadis. J. Am. Chem. Soc.,
124:14601–14607, 2002.
[177] F. Lottspeich and H. Zorbas. Bioanalytik. Spektrum, Akad. Verl., 1998.
[178] R. Winter and F. Noll. Methoden der Biophysikalischen Chemie. B.G. Teubner,
1998.
176
14. Acknowledgements
Foremost I would like to thank all those who have contributed to this work, inspired me and
supported me throughout the years of my PhD thesis.
I am grateful to my advisor Prof. Dr. Martina Havenith for giving me the opportunity to
work on this very interesting and challenging research project. Thank you for your guidance
and all kinds of support.
Prof. Dr. Christof Wöll I would like to thank for taking over the part of the co-referee. In
addition I am grateful for supporting me with the biotinylated alkylthiol and for the access to
his FTIR spectrometer.
Profound thanks go to Prof. Dr. Giacinto Scoles for giving me the opportunity to work in his
group and learn a lot about nanografting. I would also like to thank Dr. Loredana Casalis and
all members of the Scoles-group in Trieste.
I would like to thank Prof. Dr. Nils Metzler-Nolte, Dr. Ulrich Schatzschneider and Dr. Harmel
Peindy for synthesizing and providing the cymantrene labeled peptide.
My thanks go to Prof. Dr. Andreas Terfort for the PDMS stamps and the synthesis of the
biotinylated alkylthiol.
I am especially grateful for the guidance, encouragement and support given to me by Dr. Erik
Bründermann and Dr. Gerhard Schwaab during my research. Thank you also for carefully
reading this manuscript.
I would further like to thank all present and past members in the Physical Chemistry II group
for the nice working atmosphere. Special thanks go to all members of the microscopy group.
Especially to Marlena Filimon, Meike Mischo, and Dr. Jean-Sebastien Samson with whom I
shared the office. Thanks go also to Dr. Götz Wollny for the help when repairing the CO-laser.
Special thanks go to Anna Gutberlet who helped me whenever I had problems with the COlaser.
The secretaries Ursula Knieper and Sabine Weiß I would like to thank for the assistance of the
administrative questions.
I am very grateful for all the help of the technical staff, especially Reinhard Renzewitz, for
many technical short-time solutions, and Christian Fester for the fast help whenever electronic
problems occur. Thanks also to the staff of ”Glaswerkstatt, Feinmechanik, Schlosserei and
Tischlerei”.
I acknowledge the financial support from the DAAD enabling my research visit at ELETTRA
in Trieste, Italy.
I would like to express special thanks to my partner Christian Grunwald for supporting and
motivating me in the course of my doctoral study and for his faith in me.
Finally and most importantly my very special thanks and gratitude to my family - my mother,
Margarete Kopf, my sister and her husband, Marina Priester and Stefan Priester and my lovely
niece, Kathrin Priester - for all their love and support.
177
Chapter 14. Acknowledgements
178
A. Table CO laser lines
transition
transition
wavenumber
[cm−1 ]
µm-screw position
[mm]
18P17
18P16
18P15
18P14
18P13
18P12
17P18
18P11
17P17
18P10
17P16
18P9
17P15
17P13
17P12
16P18
17P11
16P17
17P9
17P8
16P14
16P12
15P18
16P11
15P17
16P12
15P16
15P14
P(17)18
P(16)18
P(15)18
P(14)18
P(13)18
P(12)18
P(18)17
P(11)18
P(17)17
P(10)18
P(16)17
P(9)18
P(15)17
P(13)17
P(12)17
P(18)16
P(11)17
P(17)16
P(9)17
P(8)17
P(14)16
P(12)16
P(18)15
P(11)16
P(17)15
P(16)12
P(16)15
P(14)15
1618
1622
1626
1629
1633
1637
1639
1640
1643
1644
1647
1647
1650
1658
1661
1664
1665
1667
1672
1676
1679
1686
1688
1690
1692
24.825
24.548
24.267
23.99
23.705
23.427
1996
1704
23.128
22.837
22.565
22.323
21.753
21.49
21.128
21.02
20.687
20.435
20.198
19.653
19.382
19.222
19.128
18.397
Continued on next page
179
Chapter A. Table CO laser lines
Table A.1 – continued from previous page
transition
transition
wavenumber
[cm−1 ]
µm-screw position
[mm]
15P13
14P19
15P12
14P18
15P10
14P16
15P9
14P15
15P8
14P14
15P7
14P13
14P12
13P18
14P11
13P7
14P10
13P16
14P9
13P15
13P14
13P13
12P19
12P18
13P11
12P17
13P10
12P15
12P14
11P20
12P13
11P19
12P12
11P18
12P11
P(13)15
P(19)14
P(12)19
P(18)14
P(10)15
P(16)14
P(9)15
P(15)14
P(8)15
P(14)14
P(7)15
P(13)14
P(12)14
P(18)13
P(11)14
P(7)13
P(10)14
P(16)13
P(9)14
P(15)13
P(14)13
P(13)13
P(19)12
P(18)12
P(11)13
P(17)12
P(10)13
P(15)12
P(14)12
P(20)11
P(13)12
P(19)11
P(12)12
P(18)11
P(11)12
1707
1709
1711
1713
1718
1721
1722
1725
1726
1729
1726
18.15
1736
1738
1740
17.892
17.153
16.972
16.702
16.455
16.202
15.81
1744
1746
1747
1750
1754
1757
1759
1763
1765
1767
1769
1775
1779
1780
1783
1784
1786
1788
1790
15.562
15.288
15.052
14.802
14.343
14.104
13.733
13.498
13.252
13.02
12.785
180
transition
transition
wavenumber
[cm−1 ]
µm-screw position
[mm]
12P10
11P16
12P9
11P15
12P8
11P14
11P13
10P19
11P12
10P18
11P11
10P17
11P10
10P16
11P9
10P14
10P13
10P12
9P18
10P11
9P17
10P10
9P16
10P9
9P15
10P8
9P14
8P20
9P13
8P19
9P12
8P18
9P11
9P10
8P16
P(10)12
P(16)11
P(9)12
P(15)11
P(8)12
P(14)11
P(13)11
P(19)10
P(12)11
P(18)10
P(11)11
P(17)10
P(10)11
P(16)10
P(9)11
P(14)10
P(13)10
P(12)10
P(18)9
P(11)10
P(17)9
P(10)10
P(16)9
P(9)10
P(15)9
P(8)10
P(14)9
P(20)8
P(13)9
P(19)8
P(12)9
P(18)8
P(11)9
P(10)9
P(16)8
1794
1796
1798
1800
1801
1804
1808
1809
1812
1813
1815
1817
1819
1821
1823
1829
1833
1837
1838
1841
1842
1845
1846
1848
1850
1852
1854
1855
1858
1859
1862
1863
1866
1870
1872
12.573
12.467
12.23
12.00
11.757
11.52
11.305
10.995
10.895
10.528
10.305
10.095
9.908
9.575
9.382
9.155
8.958
8.735
8.515
8.24
181
Chapter A. Table CO laser lines
transition
transition
wavenumber
[cm−1 ]
9P9
8P15
9P8
8P14
8P13
7P19
8P12
7P18
8P11
7P17
8P10
7P16
8P9
7P15
8P8
7P14
7P13
7P12
7P11
6P17
7P10
7P9
6P15
6P14
6P13
6P12
5P18
6P11
5P17
5P16
5P15
P(9)9
P(15)8
P(8)9
P(14)8
P(13)8
P(19)7
P(12)8
P(18)7
P(11)8
P(17)7
P(10)8
P(16)7
P(9)8
P(15)7
P(8)8
P(14)7
P(13)7
P(12)7
P(11)7
P(17)6
P(10)7
P(9)7
P(15)6
P(14)6
P(13)6
P(12)6
P(18)5
P(11)6
P(17)5
P(16)5
P(15)5
1874
1876
1878
1880
1884
1885
1888
1889
1892
1893
1896
1897
1900
1901
1903
1905
1909
1913
1917
1918
1921
1925
1927
1931
1935
1939
1940
1943
1944
1948
1952
µm-screw position
[mm]
Table A.1.: CO-laser transitions
182
8.012
7.8
7.597
7.42
7.145
6.948
6.718
6.517
6.316
6.13
5.892
5.742
5.472
5.262
5.075
4.903
4.673
4.468
4.258
B. Typical vibrational bands of
biomolecules
wavenumber
[cm−1 ]
3400
3300-3250
functional group
∼3010
∼2956
O-H stretch
amide A
100% N-H stretching
amide B
100% N-H stretching in resonance
with 1. overtone of amide II band
C-H stretch antisym.
in N+ -(CH3 )3
C-H stretching antisym. in -CH=CH- (cis)
C-H stretching antisym. in CH3
∼2921
C-H stretching antisym. in CH2
∼2872
C-H stretching sym. in CH3
∼2850
C-H stretching sym. in CH2
1745-1735
1650
C=O stretch (ester)
O-H scissoring
C=C (cis) stretching
amide I
80% C=O stretching
loops
beta sheet
alpha helix
disordered structures
beta sheet
amide II
60% N-H bending
40% C-N stretching
∼3100
3028-3050
1600-1700
1690-1660
1680-1670
1658-1650
1650-1640
1640-1620
1575-1480
molecules
water
proteins
proteins
phosphatidylcholine
unsaturated lipids
lipids, proteins, nucleic
carbon hydrates
carbon hydrates
carbon hydrates
carbon hydrates
lipids, thymine, uracile
water
unsaturated lipids
proteins
acids,
acids,
acids,
acids,
proteins
183
Chapter B. Typical vibrational bands of biomolecules
∼ 1515
1490
∼ 1465
∼ 1450
1405
∼ 1400
∼ 1380
1330-1230
1330-1180
1250-1220
1200-900
∼ 1170
∼ 1090
∼1070
∼1050
∼970
970
∼725
∼720
625-770
tyrosine (amino acid side chain)
C-H deformation in N+ -(CH3 )3
C-H deformation in CH2
C-H deformation antisym. in CH3
C-H deformation sym. in N+ -(CH3 )3
C=O stretch in COO−
C-H deformation sym. in CH3
amide III
40% C-N stretching
30% N-H bending
20% C-C stretching
C-H wagging in CH2
P=O stretch antisym. in PO−
2
C-O-P und C-O-C stretch and ring
CO-O-C stretch
P=O stretch sym. in PO−
2
CO-O-C stretch sym.
different overlapping bands from
C-O stretch in CH2 OH,
C-O stretch and C-O bending in C-OH,
C-O-P stretch
ribose
C-N+ -C stretch antisym. in N+ -(CH3 )3
amide V
N-H bending
C-N torsion
C-H rocking in CH2
amide IV
40% C=O bending
39% C-C stretching
proteins
phosphatidylcholine
lipids, proteins, nucleic acids
lipids, proteins
phosphatidylcholine
fatty acids, amino acid side chains
lipids, proteins
proteins
nucleic acids, lipids
carbon hydrates, cholesterol, ester
lipids
nucleic acids,
lipids
ribose scaffold of nucleic acids
carbon hydrates, lipids
RNA
phosphatidylcholine
proteins
proteins
proteins
Typical infrared vibrational modes of biomolecules and water [177, 178]
184
C. Purchasing of synthetic
polynucleotides
Nowadays many companies, like Biomers, Thermo electron or Sigma Aldrich, purchase
DNA with desired sequences and modifications such as fluorophores or specific linkers.
When ordering DNA one has to pay attention to some details:
• Synthesis scale In general the synthesis scale refers to the amount of solid-phase
support carrying the start base that is used for the synthesis. It does not refer to
the final amount of the oligonucleotide that will be delivered!
• Quantification of oligonucleotides The quantity of an oligonucleotide is usually
given as its total optical density (OD value), as amount of substance (nmol) or as
its available mass (µg). In practice the OD value is measured experimentally and
the other values are calculated. The OD value of a sample at a wavelength of 260
nm is defined as the extinction that occurs in measuring absorption of the sample
in 1 ml aqueous solution in a 1 cm cuvette. According to the Lambert-Beer-law
(E = ε*c*d) one can calculate from the extinction E (OD value) the concentration
c and from this the amount of substance. The extinction coefficient is different
for each oligonucleotide sequence and can be predicted computationally using so
called nearest-neighbour methods.
• Purification grades Desalted means that excess salts and other lower molecular
weight compounds are removed. The oligonucleotide solution contains besides the
desired full-length oligonucleotide also all incomplete chains which form during
synthesis. After desalting process the incomplete chains remain in the solution.
High pressure liquid chromatography (HPLC) purification is recommended when
a highly purified DNA with a length of up to 80 bases is desired. Typical obtained
purity is 95-99%.
Polyacrylamide gel electrophoresis (PAGE) purification is mainly used for very
long polynucleotides and DNA used for cloning applications.
• Storage Oligonucleotides are best stored in dry state. At room temperature dry
oligonucleotides are stable several days, at 4◦C for several weeks and at -20◦C for
several month. Dissolving immediately before use is best and repeated freeze/thaw
cycles should be avoided. To this end aliquoting of stock solutions is recommended.
Some companies offer the possibility to deliver dried aliquots of a desired amount.
185
Chapter C. Purchasing of synthetic polynucleotides
• Dissolving Since the oligonucleotide pellets can stick at the cap of the tube the
tube should be shortly centrifuged before opening. Oligonucleotides can be dissolved in sterile water at neutral pH or in buffer at a slightly basic range (e.g.
Tris-HCl, TE (10mM Tris, 1 mM EDTA = 1xTE), PBS or TSE (25mM Tris,
100mM NaCl, 0.1 mM EDTA = 1xTSE). When using water from purification
systems one has to check the pH because such water is often slighly acidic!
186
D. Symbols and Abbreviations
symbol
abbreviation
Å
α
∆G
∆H
ε
λ
µCP
ν
ν̃
σ
τ
a
A
AFM
Au
BAT
BaF2
bp
C
CaF2
CO
CO2
CymPntCys
DFT
meaning
Ångstrom
polarizability
Gibbs free energy
enthalpy
dielectric constant
wavelength
microcontact printing
frequency
wavenumber
scattering cross-section
time constant
AFM tip curvature radius
adenine
atomic force microscope,
atomic force microscopy
gold
biotinylated alkylthiol
barium fluoride
base pair
cytosine
calcium fluoride
carbon monoxide
carbon dioxide
cymantrene labeled peptide
density functional theory
187
Chapter D. Symbols and Abbreviations
Table
symbol
abbreviation
DNA
ds
dsDNA
DTGS
EDTA
EG
EtOH
f
FTIR
FZ curve
GIR
G
H2 O2
H2 SO4
IR
IRRAS
KBr
MCH
MCT
n
ODT
OEG
OPO
PDMS
PMMA
PVD
RIU
SAM
SEM
s-SNIM
188
D.1 – continued from previous page
meaning
deoxyribonucleic acid
double-stranded
double-stranded DNA
deuterated triglycine sulfate
ethylenediaminetetraacetic acid
ethylene glycole
ethanol
frequency
fourier transform infrared
force distance curve
grazing incident reflection
guanine
hydrogen peroxide
sulfuric acid
infrared
infrared reflection absorption spectroscopy
potassium bromide
6-mercapto-1-hexanol
mercury cadmium telluride
refractive index
1-octadecanethiol
oligo (ethylene glycole)
opto parametric oscillator
poly(dimethylsiloxan)
poly(methylmetacrylate)
physical vapor deposition
refractive index units
self-assembled monolayer
scanning electron microscope
scanning electron microscopy
scattering scanning near-field infrared microscopy
scattering scanning near-field infrared microscope
Table D.1 – continued from previous page
symbol
meaning
abbreviation
SNIM
scanning near-field infrared microscope
scanning near-field infrared microscopy
SNOM
scanning near-field optical microscope
scanning near-field optical microscopy
SNR
signal-to-noise ratio
SPL
scanning probe lithography
SPM
scanning probe microscope
scanning probe microscopy
SPR
surface plasmon resonance
ssDNA
single-stranded DNA
STE buffer
1 M NaCl TE buffer
Tm
melting temperature
T
thymine
TE buffer
10 mM Tris buffer pH 7.2, 1 mM EDTA
TSG
template stripped gold
UHV
ultra high vacuum
UV
ultra violet
V
volt
VIS
visible
189
191
Chapter E. Curriculum Vitae and Publications
E. Curriculum Vitae and Publications
Personal details
Name
Date of Birth
Place of Birth
Marial status
Ilona Kopf
24.08.1977
Remscheid, Germany
not married
Education
10/2004 - 01/2009
Doctorate Programme at the Graduate School of Chemistry and
Biochemistry
Department of Physical Chemistry II (Prof. M. Havenith)
Ruhr-University Bochum, Germany
Title of Dissertation:”Near-field infrared microscopy
applied to laterally structured self-assembled monolayers”
minor subject: ”Photonics” and ”Optoelectronic devices”
10/1999 - 08/2004
Diploma degree in Biochemistry
Faculty for Chemistry and Biochemistry
Ruhr-University Bochum, Germany
02/2004 - 08/2004 Diploma thesis
Department of Physical Chemistry II (Prof. M. Havenith)
Title of Diploma thesis:”Nahinfrarot-Spektroskopie von Wasser in
lebenden Zellen und Zellgewebe”
1996 - 1999
Ricarda-Huch-Schule, Hannover, Germany
June, 1999 ”Abitur” (german university entrance qualification)
1994 - 1995
Ortlinghause-Werke, Wermelskirchen, Germany
Industrial business management assistant
June, 1995 job training aborted
1988 - 1994
Städtisches Gertrud-Bäumer Gymnasium, Remscheid, Germany
June, 1994 Fachoberschulreife (Certificate of Secondary Education)
Städtische Gemeinschaftsgrundschule Hasten, Remscheid, Germany
1984 - 1988
192
Scholarships &
Foreign studies
short time scholarship of Deutschen Akademischen
Austauschdienstes e.V. (DAAD)
09/2007 & 10/2007, Nanostructure Laboratory (Prof. G. Scoles)
ELETTRA Sincrotrone Trieste S.C.p.A., Trieste, Italy
Travel grants
02/2008 Ruth und Gerd Massenbergstiftung, Winter College Trieste
Schools/Colleges
09/2005, Summer School: Physics of Imaging, Bad Honnef, Germany
02/2008, Winter College on micro and nano photonics for life sciences
Trieste, Italy
Publications
• Ilona Kopf, Christian Grunwald, Erik Bründermann, Loredana Casalis, Giacinto Scoles,
Martina Havenith:
Detection of hybridization on nanografted oligonucleotides using scanning near-field infrared microscopy,
in preparation
• Erik Bründermann, Ilona Kopf, Martina Havenith:
Chemical nanoscopy of cell-like membranes,
Proceedings of SPIE, 7188 (2009), submitted, Invited Paper
• Ilona Kopf, Jean-Sebastien Samson, Götz Wollny, Christian Grunwald, Erik Bründermann, Martina Havenith:
Chemical Imaging of microstructured self-assembled monolayers with nanometer resolution,
J. Phys. Chem. C 11(23), S.8166-8171 (2007)
• Erik Bründermann, Andreas Bergner, Frank Petrat, Robert Schiwon, Götz Wollny, Ilona
Kopf, Herbert de Groot, Martina Havenith:
Fast quantification of water in single living cells by near-infrared microscopy,
Analyst 129, 893 (2004)
Talks at international and national conferences
• Winter College on micro and nano photonics for life sciences, 2008, Trieste, Italy
I. Kopf, J.-S. Samson, G. Wollny, Ch. Grunwald, E. Bründermann, M. Havenith:
Scanning near-field infrared microscopy applied to self-assembled monolayers
• SLONANO, 2007, Ljubljana, Slovenia
Infrared nanoscopy of structured SAMs
• Frühjahrestagung der Deutschen Physikalischen Gesellschaft, 2005, Berlin, Talk
I. Kopf, A. Bergner, E. Bründermann, F. Petrat, M. Havenith:
Quantifizierung von Wasser in lebenden Zellen durch Nahinfrarot-Mikroskopie
193
Chapter E. Curriculum Vitae and Publications
Posters at international and national conferences
• 107. Bunsentagung, 2008, Saarbrücken, Poster
I. Kopf, Ch. Grunwald, E. Bründermann, L. Casalis, G. Scoles, M. Havenith:
Hybridization detection at the nanoscale using scanning near-field infrared microscopy
selected to be a HOT TOPIC poster among 9 others out of 264 posters
• 106. Bunsentagung, 2007, Graz, Austria, Poster
Chemical imaging of microstructured SAMs with nanometer resolution
• 5. Deutsches BioSensor Symposium, 2007, Bochum, Poster
I. Kopf, G. Wollny, J.-S. Samson, E. Bründermann, M. Havenith:
Labelfree infrared nanoscopy
poster prize
• Frühjahrestagung der Deutschen Physikalischen Gesellschaft, 2006, Frankfurt am Main,
Poster
I. Kopf, G. Wollny, J.-S. Samson, E. Bründermann, M. Havenith:
Highly sensitive absorption contrast imaging with a near-field infrared nanoscope
Television contributions
• Magazin Hitec 3sat/ZDF: Vorstoß ins Unsichtbare, 13.06.2004
short summary about near-infrared microscopy
International and national conference contributions as co-author
• Materialwissenschaftlicher Tag, 2008, Bochum, Poster
E. Bründermann, I. Kopf, F. Ballout, M. Filimon, O.̈ Birer, D. Schmidt, M. Havenith:
Development of a scanning near field infrared microscope - chemical mapping on a nm
scale
• Winter College on micro and nano photonics for life sciences, 2008, Trieste, Italy, Poster
J. Mondry, M. Mischo, I. Kopf, M. Filimon, E. Bründermann, M. Havenith:
Labelfree investigation of drug dependent water dynamics in single living cells by nearinfrared microscopy
• Winter College on micro and nano photonics for life sciences, 2008, Trieste, Italy, Poster
A. Kress, J.-S. Samson, I. Kopf, F. Ballout, D. Schmidt, E. Bründermann, G. Schwaab,
M. Havenith:
Imaging of Human Hair with scanning near-field infrared microscopy (SNIM) and confocal Raman microscopy
• BMT, 2006, Zürich, Talk
M. P. Mienkina, K. Hensel, T. N. Le, N. C. Gerhardt, I. Kopf, E. Bründermann, M.
Havenith, M. Hofmann, G. Schmitz:
Experimentelle Charakterisierung von Ferucarbotran als photoakustisches Kontrastmittel
194
• 104. Bunsentagung, 2005, Frankfurt am Main, Poster
E. Bründermann, A. Bergner, I. Kopf, G. Wollny, F. Petrat, M. Havenith:
Fast quantification of unlabeled molecules in living cells by infrared laser microscopy
• Frühjahrestagung der Deutschen Physikalischen Gesellschaft, 2004, München, Poster
A. Bergner, E. Bründermann, I. Kopf, R. Schiwon, M. Havenith:
Near infrared microspectroscopy of biomolecules
• RISBM, 2004, Jena, Poster
A. Bergner, E. Bründermann, I. Kopf, R. Schiwon, M. Havenith:
Near infrared Microspectroscopy of Biomolecules
195

Near-field infrared nanoscopy applied to laterally structured self

Transcription

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