We thank M. Hempenius and E. Benetti for fruitful discussions, Y. Yu for careful checking of the recipe, M. Vlot for the image design of Figure 1, C. Padberg and K. Smit for technical support. E.K. acknowledges the Netherlands Organization for Scientific Research (NWO, TOP Grant 700.56.322, Macromolecular Nanotechnology with Stimulus Responsive Polymers) for financial support. SdB has been supported by the Foundation for Fundamental research on Matter (FOM), which is financially supported by the Netherlands Organization for Scientific Research (NWO).

NOTE: Figure 2 illustrates the sample preparation procedure. Polymer brushes were grafted from silicon (Si) substrates (path (a)), from gold-coated substrates (100 nm gold evaporated on Si wafer having a 10 nm Cr adhesion layer, path (b)), and from gold colloidal AFM probes (6 &#956;m diameter, path (c)) by surface initiated atom-transfer radical polymerization (SI-ATRP)24. The AFM measurements were performed on a Multimode AFM with the low-noise head, vertical engage scanner and a liquid cell.
1. Sample Preparation
<ol>
	<li>Substrate preparation
	<ol>
		<li>Initiator deposition on silicon surfaces.
		<ol>
			<li>Clean the silicon substrates with chloroform, then piranha solution, subsequently rinse with purified water, ethanol and chloroform.</li>
			<li>Place the dry substrates into a desiccator around a vial containing 50 &#181;l (3-aminopropyl) triethoxysilane.</li>
			<li>Evacuate the desiccator with a rotary vane pump for 15 min and close it subsequently. Allow the vapor deposition to proceed O/N.</li>
			<li>Prepare a solution of 40 ml toluene (degased) and 40 &#181;l triethylamine in an Erlenmeyer flask.</li>
			<li>Place the substrates in the flask and drop-wise add 40 &#181;l 2-bromo-2-methylpropionyl. Keep the substrates in the solution, to avoid deposition of salt crystals formed in solution.</li>
			<li>Stir the solution for 4 hr.</li>
			<li>Rinse the substrates with toluene and ethanol. Next, place the substrates into the polymerization vials.</li>
		</ol>
		</li>
		<li>Initiator deposition on gold surfaces
		<ol>
			<li>Prepare a monolayer solution: dissolve 2-bromo-2-methyl-propionic acid 11-[11-(2-bromo-2-methyl-propionyloxy)-undecyldisulfanyl]-undecyl ester in 20 ml degassed chloroform (0.2 mM).</li>
			<li>Transfer ca. 1.5 ml monolayer solution into a small vial that was previously flushed with Argon.</li>
			<li>Clean the gold-coated substrates with chloroform and piranha solution. Subsequently rinse with purified water, ethanol and chloroform.</li>
			<li>Immerse the cleaned substrates into the monolayer-solution and close the flask. Store O/N in a dark place.</li>
			<li>Clean the gold colloidal probes with ethanol and chloroform by dipping them into the solvents.</li>
			<li>Immerse the colloidal probes into the vial containing the monolayer solution. Close firmly and keep it in a dark place O/N.</li>
			<li>Remove the substrates from the solution, wash with chloroform and ethanol. Next, transfer the initiator coated substrates into the polymerization flask.</li>
			<li>Remove the colloidal probes from the monolayer solution, wash with chloroform and ethanol. Next, transfer each probe separately into the polymerization vial.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Polymerization
	<ol>
		<li>SI-ATRP of PMMA
		<ol>
			<li>Purge the flask and vials, containing the initiator-covered substrates (both Si and gold) and the colloidal probes with Argon for 30 min.</li>
			<li>Dissolve 10 g methyl methacrylate (MMA) in the ATRP medium (10 ml methanol/water mixture with ratio 5:1) and degas the solution for 2 hr.</li>
			<li>Add 145 mg CuBr and 320 mg 2,2-bipyridine into a flask equipped with a magnetic stirring bar, and deoxygenize by 3 vacuum-Ar backfill cycles.</li>
			<li>Transfer the degassed monomer solution into the flask (containing the copper) and stir for another 15 min until a clear brown solution is observed.</li>
			<li>Attach a needle to a 2 ml syringe and flush the needle and the syringe with argon 2-3 times.</li>
			<li>Withdraw 1 ml polymerization solution with the syringe and inject the content into the small reaction vial, containing the colloid probe.</li>
			<li>Withdraw the remaining solution with the syringe and inject the content into the reaction vial, containing the flat substrates. Add enough solution to submerge each sample completely.</li>
			<li>Conduct the polymerization for 40 hr at RT.</li>
			<li>Remove the samples from the polymerization solution and wash with ethanol and chloroform in multiple cycles. The coloring of the sample indicates the presence of the brush.</li>
			<li>Finally, dry the substrates under a stream of nitrogen.</li>
			<li>Keep flat substrates in a nitrogen box. Store polymer brush modified probes in toluene.</li>
		</ol>
		</li>
		<li>SI-ATRP of PNIPAM10
		<ol>
			<li>Purge the flask and vials, containing the initiator-covered substrates (both Si and gold) and the colloid probes with argon for 30 min.</li>
			<li>Dissolve 5.6 g N-isopropylacrylamide and 320 &#181;l PMDETA in the ATRP medium (1.6 ml water and 18 ml methanol) and degas the solution for 2 hr.</li>
			<li>Add 76 mg CuBr into a flask equipped with a magnetic stirring bar, and deoxygenize by 3 vacuum-Ar backfill cycles.</li>
			<li>Transfer the degassed monomer solution into the flask that contains the copper. Stir for another 15 min until a clear green solution is observed.</li>
			<li>Attach a needle to a 2 ml syringe and flush the needle and the syringe with argon 2-3 times.</li>
			<li>Withdraw 1 ml polymerization solution with the syringe and inject the content into the small reaction vial, containing the colloid probe. Repeat for all probes.</li>
			<li>Withdraw the remaining solution with the syringe and inject the content into the reaction flask, containing the flat substrates. Add enough solution to submerge each sample completely.</li>
			<li>Conduct the polymerization for 2 hr at RT.</li>
			<li>Remove the samples from the polymerization solution and wash with ethanol and water through multiple cycles. The appearing color of the sample indicates the presence of the brush.</li>
			<li>Immerse the sample into 0.1 M EDTA solution and keep it in the solution O/N to remove all the copper. Wash with purified water and ethanol.</li>
			<li>Keep the flat substrates in a nitrogen box. Store the PNIPAM brush modified probes in purified water.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Brush characterization, Fourier Transform Infrared (Figure&#160;3) 
	NOTE: Use grazing-angle FTIR mode for polymer brushes on gold substrate and transmission FTIR mode for polymer brushes on Si substrate. Dry all samples carefully. Any water and solvent contamination can cause serious damage of the detector.&#160;
	<ol>
		<li>Start up the equipment according to the manual provided by the manufacturer.</li>
		<li>Set the following parameters for sample measurement: range between 900 and 3,700 cm-1 with a resolution of 4 cm-1 and average 32 scans in each measurement.&#160;</li>
		<li>Clean a blank substrate according to the protocol used for sample preparation, dry carefully and place it to the measurement chamber.</li>
		<li>Apply vacuum for 4 hr. Next, record a background scan.</li>
		<li>Remove the blank sample and place the dry polymer brush coated sample in the sample chamber.</li>
		<li>Apply vacuum in the sample chamber and record a sample-scan every 30 min for 5 hr.</li>
		<li>Identify all peaks in the spectra to confirm chemical composition of the polymer brush.&#160;(Only analyze the scans with a straight baseline.)</li>
	</ol>
	</li>
</ol>
2. AFM Measurement
<ol>
	<li>Scratch the PMMA brush covered substrate carefully with a needle, rinse with a good solvent to remove the excess of polymer and dry it.</li>
	<li>Mount the samples into the AFM instrument and mount the probe into the liquid cell.</li>
	<li>Align the laser on the end of the cantilever.</li>
	<li>With use of the camera, align the tip above a scratch.</li>
	<li>Before approaching and engaging the tip, set the scan size 0 nm. Next, engage the tip to the surface.</li>
	<li>Go to &#8216;ramp mode&#8217; and determine the deflection sensitivity via force distance curves. Calibrate the normal spring constant using the &#8216;thermal tune&#8217; as implemented in the software and torsional spring constant of the cantilever using the method of Wagner et al.25
	<ol>
		<li>Capture the torsional thermal noise of the cantilever in air for 2 sec using the high speed data capture in the software (6.25 MHz).</li>
		<li>Convert the thermal noise into the power spectral density (V2/Hz) using the Fourier Transform.</li>
		<li>Determine the fundamental resonance and the quality factor of the cantilever in air using the resonance peak in the power spectral density and the equation for a simple harmonic oscillator including baseline noise (eq. 2 of Ref. 25).</li>
		<li>Calculate the torsional spring constant using the inplane dimensions of the cantilever (length and width), the density and viscosity of the surrounding medium (air) and the quality factor and resonance frequency determined in step 2.6.3 using the method of Sader26. 
		NOTE: We used the tool provided on the website of John Sader: www.ampc.ms.unimelb.edu.au/afm/calibration.html.</li>
		<li>Calculate the torsional angle deflection sensitivity of the cantilever using eq. 6 and 7 of Ref. 25.</li>
		<li>Convert the torsional spring constant and deflection sensitivity into the lateral spring constant and deflection sensitivity using the size of the colloid, the thickness of the cantilever and eq. 8 of Ref. 25. 
		NOTE: the detector signal can now be converted into a force via: force [N] = lateral spring constant [N/m] * lateral deflection sensitivity [m/V] * detector signal [V].</li>
	</ol>
	</li>
	<li>Measure the brush height of the dry brush by imaging the brush at a scratch at the lowest possible deflection setpoint. Determine the brush height from the line-scans of the captured image.</li>
	<li>Solvate the brush in acetophenone by gently applying it to the surface with a syringe. 
	NOTE: The color of the sample changes as the solvent evaporates. This allows for following the drying process.</li>
	<li>Mount the sample to the AFM. Align the vertical laser signal to -1.0 V. Set the deflection setpoint to 0 V and engage the cantilever and surface. 
	NOTE: After bringing the cantilever in contact with the surface, the acetophenone moves into the brush on the colloid creating a capillary bridge between the tip and the surface.</li>
	<li>Set the scan size to 40 &#956;m, disable the slow scan axes and set image aspect ratio to 1:4. Record the height and the friction image channels (both trace and retrace).</li>
	<li>When the images are captured withdraw the cantilever.</li>
	<li>Apply a drop of water on the PNIPAM brush-covered surface, to solvate the PNIPAM brush.</li>
	<li>Lift the head and quickly replace the PMMA surface by the solvated PNIPAM surface to create the immiscible system. Be quick when exchanging the surfaces, to avoid evaporation of the acetophenone from the PMMA brush on the colloidal probe.</li>
	<li>Engage the tip and surface, and record images with same parameters as previously.</li>
</ol>

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The authors have nothing to disclose.

The presented results show that the friction, for immiscible systems of individually solvated brushes, is strongly reduced compared to traditional miscible systems of two of the same solvated brushes. The preferred absorbance of the different solvents in the two brushes prevents the brushes from interdigitating and consequently a major source of wear and dissipation in polymer brush friction is eliminated. The presented method is therefore fundamentally different from sliding dry hydrophilic on hydrophobic brushes, where the friction will be determined by brush-specific interactions27. In fact, upon shearing PMMA on PNIPAM (collapsed height 166 nm) without solvents, we found that the friction was 50% higher compared to dry PMMA on PMMA (collapsed height 236 m).
As already pointed out shortly in the notes of the &#8216;Protocol&#8217; section, there are a couple of crucial points that need to be kept in mind while performing these particular experiments: Firstly, acetophenone is a better solvent for PNIPAM than water. Thus, care should be taken that acetophenone does not enter the PNIPAM brush by wetting the PNIPAM brush with plenty of water. Since acetophenone and water do not mix, the acetophenone will now not enter the PNIPAM brush. That is why we did not immerse our system completely in acetophenone, but instead created an acetophenone capillary for the miscible system. Another reason for incomplete immersion is that complete immersion results in too strong hydrodynamics, such that we only measured the Stokes drag on the colloid and cantilever. Secondly, in AFM experiments the torsional and normal spring constants are coupled. Cantilevers with a low normal spring constant will also have a relatively low torsional spring constant and vice versa. This limits the lowest measurable friction coefficient to &#62;10-3. Thus, in order to measure the full friction reduction, the friction for the miscible system needs to be high. This is achieved by using long high-density brushes and a relatively high shear velocity of typically 100 m/sec. Moreover, the capillary between the brushes also increases the friction forces. We measured the lowest friction coefficient, for an immiscible system22, of &#956; = 0.003 under an estimated normal stress of 200 kPa. Using the same experimental conditions, we found that &#956; = 0.15 for the miscible system.
Note that the experiments were performed in a controlled laboratory environment and that surfaces used in industry are not as ideal as used in the presented experiments. Most surfaces have a non-uniform roughness distribution28 and thus many asperities of different shapes and sizes. During the collision of two brush-bearing asperities, the friction is composed out of different dissipation channels29. Next to steady-state dissipation mechanisms, such as interdigitation and solvent flow, there will be hysteretic effects in the shape30 due to the slow relaxation time of the polymers and solvent. Besides, capillaries are formed and broken. In the traditionally used miscible brush systems, transient interdigitation31 amplifies shape- and capillary hysteresis. With the immiscible system presented here, transient interdigitation is eliminated too. Moreover, capillary hysteresis can be circumvented by application of two immiscible solvents. Therefore, also for the more common rough surfaces, friction and wear will be reduced using immiscible brush systems22. The main source of friction that remains is brush deformation. Anchoring polyzwitterionic polymers, which are known for their intrinsic low friction32, onto one of the surfaces can minimize the latter. In such systems the osmotic pressure of the solvent is high resulting in little brush deformation under high normal loads.
The presented method of immiscible brush systems can be applied in almost any system where low friction is desirable. The method functions well under high pressures. However, care should be taken that the temperature is kept around RT. High temperatures damage the polymers, which will cause liquid-flow out of the contact and consequently high friction. Examples of potential application are: syringes, piston systems, axle bearings and hinges.

Perfect lubricants reduce friction and wear for solids in relative motion even when normal loads are high. To achieve this, the lubricant should remain in the contact during sliding and at rest. However, under a positive normal load, simple, low-viscosity liquids are quickly squeezed out of the contact area and even higher-viscosity oils are expelled eventually. Yet, biological contacts, e.g., in human joints, remain lubricated with low-viscosity fluids at all times. Nature realizes such efficient lubrication using sugar chains attached to solid surfaces1. The hydrophilic sugar chains keep an aqueous liquid in the contact provided that the normal pressure does not exceed the osmotic pressure of the solvent2. Therefore, a lot of effort has been directed towards mimicking biological lubricants by grafting polymers to solid surfaces forming so-called polymer brushes3-12.
When two opposing polymer brushes are brought into contact, segments of the polymer chains on one side can move into the brush chain segments at the opposite side. This effect is called interdigitation13. When the brushes are in relative sliding motion, interdigitation is the main source of wear14 and friction15-17. In fact, recently, friction-velocity relations for sliding polymer brushes have been derived18. These scaling laws are based on interdigitation and the consequent stretching and bending of the polymers upon sliding. The main characteristics agree with results of surface forces apparatus experiments19 and of molecular dynamics (MD) simulations20. In the latter the degree of overlap can be directly quantified. Moreover, it was shown that the overlap between polyelectrolyte brushes can be tuned by applying an electric field21. Thus, if interdigitation can be circumvented, friction and wear in these systems would be significantly reduced.
In a recent publication22 we have shown via MD simulations that two immiscible solvated polymer brush systems prevent overlap between the brushes. Moreover, upon sliding the brushes, we found a decrease of the friction force by two orders of magnitude compared to the traditional miscible brush systems, in excellent agreement with our atomic force microscopy (AFM) measurements. Here, we explain in detail how to set up the AFM experiments of Ref. 22. The basic principle is sketched in Figure&#160;1. On the two counter-surfaces, two different brushes, each solvated by their own preferred solvent, are needed. In this configuration each brush remains in its own solvent. Consequently, polymer segments from one brush do not penetrate into the other brush. Poly(methyl methacrylate) (PMMA) is grafted from an AFM colloidal probe and the brush is solvated by acetophenone. From the flat surface poly(N-isopropylacrylamide) (PNIPAM) is grafted and solvated in water. To compare the present system to traditional miscible systems, a second flat counter-surface bearing a PMMA brush solvated in acetophenone is made. The measured friction force upon sliding the immiscible system of PMMA on PNIPAM is approximately 1% of the friction for the miscible system of PMMA on PMMA. Note that the use of these particular brush systems is just one example. The presented method is generic and works due to the preferred absorbance of the solvents in the different brushes. Therefore, more types of brushes are expected to be applicable, as long as the chosen solvents de-mix in the two brushes. The effect is amplified by using two non-mixing solvents (like acetophenone and water) such that an extra slippery fluid-on-fluid-sliding interface is created22,23.

<table><tbody><tr><td>Methyl methacrylate</td><td>Sigma-Aldrich</td><td>M55909&amp;nbsp;</td><td>Monomer for PMMA synthesis, cleaned by pressing through a basic alumina column</td></tr><tr><td>(3-aminopropyl)triethoxysilane</td><td>Sigma-Aldrich</td><td>440140</td><td>vapor deposited silane monolayer</td></tr><tr><td>triethylamine</td><td>Sigma-Aldrich</td><td>T0886</td><td>Reagent for the ATRP initiator moiety coupling.</td></tr><tr><td>2-bromo-2-methylpropionyl</td><td>Sigma-Aldrich</td><td>252271</td><td>ATRP initiator moiety.</td></tr><tr><td>toluene</td><td>Biosolve</td><td>20150501</td><td>Coupling medium for ATRP moiety</td></tr><tr><td>CuBr</td><td>Sigma-Aldrich</td><td>212865</td><td>ATRP catalyst.</td></tr><tr><td>2,2&amp;prime;-Bipyridyl</td><td>Sigma-Aldrich</td><td>14453</td><td>Cu complexing ligand for ATRP of MMA</td></tr><tr><td>&lt;em&gt;N,N,N&amp;prime;,N&quot;,N&quot;&lt;/em&gt;-Pentamethyldiethylenetriamine</td><td>Sigma-Aldrich</td><td>369497</td><td>Cu complexing ligand for ATRP of NIPAM</td></tr><tr><td>acetic acid 98-100%</td><td>Merck</td><td>8187551000</td><td>For cleaning CuBr.</td></tr><tr><td>Sulfuric acid</td><td>Sigma-Aldrich</td><td>320501</td><td>For the preparation of Piranha solution</td></tr><tr><td>Hydrogen peroxide 33%</td><td>Merck</td><td>1.07210.1000</td><td>For the preparation of Piranha solution</td></tr><tr><td>Ethanol</td><td>Merck</td><td>1.00983.1000</td><td>For cleaning substrates.</td></tr><tr><td>Basic aluminum oxide 60&amp;nbsp;</td><td>Merck</td><td></td><td>For cleaning monomers.</td></tr><tr><td>Chloroform</td><td>Biosolve</td><td>3080501</td><td>For monolayer deposition and substrate cleaning.</td></tr><tr><td>Methanol</td><td>Biosolve</td><td>13680501</td><td>For polymerization medium.</td></tr><tr><td>Acetophenone</td><td>Acros Organics</td><td>102410010</td><td>For AFM measurement environment.</td></tr><tr><td>&lt;em&gt;N&lt;/em&gt;-isopropyl acrylamide</td><td>Acros Organics</td><td>412780250</td><td>Monomer for PNIPAM synthesis, recrystallized from toluene/hexane</td></tr><tr><td>Poly(ethylene glycol) methacrylate&amp;nbsp;</td><td>Sigma-Aldrich</td><td>409529</td><td>Monomer for Si-POEGMA synthesis, cleaned by pressing through a basic alumina column.</td></tr><tr><td>MilliQ water</td><td>MilliQ Advantage A 10 purification system&amp;nbsp;</td><td></td><td>ATRP medium, AFM measurement environment and for substrate cleaning.</td></tr><tr><td>Silicon substrates&amp;nbsp;</td><td></td><td></td><td></td></tr><tr><td>Gold coated substrates</td><td></td><td></td><td></td></tr><tr><td>AFM probe, CP-FM-Au</td><td>SQube</td><td></td><td>AFM measurement</td></tr><tr><td colspan="4">[header]</td></tr><tr><td>dithiodiundecane-11,1-diybis[2-bromo-2-methlpropanoate] (DTPR)</td><td></td><td></td><td>Initiator, for Si-ATRP on gold surfaces.&lt;br/&gt; Synthesized according to &lt;em&gt;Macromolecules&lt;/em&gt;, &lt;strong&gt;33&lt;/strong&gt;, 597-605 (2000).</td></tr><tr><td>Atomic Frorce Microscope</td><td>Bruker Multimode V controller</td><td></td><td></td></tr></tbody></table>

preparation and friction force microscopy measurements of immiscible, opposing polymer brushes

Solvated polymer brushes are well known to lubricate high-pressure contacts, because they can sustain a positive normal load while maintaining low friction at the interface. Nevertheless, these systems can be sensitive to wear due to interdigitation of the opposing brushes. In a recent publication, we have shown via molecular dynamics simulations and atomic force microscopy experiments, that using an immiscible polymer brush system terminating the substrate and the slider surfaces, respectively, can eliminate such interdigitation. As a consequence, wear in the contacts is reduced. Moreover, the friction force is two orders of magnitude lower compared to traditional miscible polymer brush systems. This newly proposed system therefore holds great potential for application in industry. Here, the methodology to construct an immiscible polymer brush system of two different brushes each solvated by their own preferred solvent is presented. The procedure how to graft poly(N-isopropylacrylamide) (PNIPAM) from a flat surface and poly(methyl methacrylate) (PMMA) from an atomic force microscopy (AFM) colloidal probe is described. PNIPAM is solvated in water and PMMA in acetophenone. Via friction force AFM measurements, it is shown that the friction for this system is indeed reduced by two orders of magnitude compared to the miscible system of PMMA on PMMA solvated in acetophenone.

Figure 4 shows representative AFM force traces upon sliding both the miscible and the immiscible polymer brush systems. The friction force F is normalized by the friction force at steady state sliding Fsym for the symmetric, miscible system. The swollen brush height in these experiments was 1,010 nm for PMMA and 532 nm for PNIPAM. The force traces are captured after following the procedure described in the Protocol section. In these experiments the surface was moved back and forth with a velocity v of 80 &#956;m/sec while applying a normal load of 30 nN. The difference in friction force for the miscible (left panel) and immiscible (right panel) brush systems can be clearly observed. The steady state friction force in the left panel is 90x higher than the steady state friction force in right panel. For the immiscible system the measured friction force is typically 0.5&#8211;2% of the friction force measured for the miscible system. Though the exact friction reduction depends on the grafting density, degree of polymerization, amount of solvent, and (weakly) on the normal load and sliding velocity, it is always around two orders of magnitude. If we increase the sliding velocity for the system described above by a factor 5 (to 400 &#956;m/sec), the friction reduction decreases by 2%. If we increase the normal load by a factor 10 (to 300 nN), the friction reduction decreases by 3%.
<img alt="Figure 1" src="/files/ftp_upload/52285/52285fig1highres.jpg" /> 
Figure 1. Schematic sketch of the setup. Left panel shows the miscible system, where the same polymers are grafted from the surface and the colloid. The brushes are solvated in a one-phase liquid. The right panel shows the immiscible system of two different polymer brushes. Each brush is solvated in its own preferred liquid. In traditional miscible systems the polymers of the opposite brushes overlap. For the immiscible system, opposite brushes do not interdigitate such that friction and wear during sliding is reduced.
<img alt="Figure 2" src="/files/ftp_upload/52285/52285fig2highres.jpg" width="700px" /> 
Figure 2. Schematic sketch of the sample preparation procedure as described in the Protocols section. From left to right shows the procedure of brush preparation via initiator deposition and surface initiated atom transfer radical polymerization (SI ATRP). Path (A) describes the brushes grafted from silicon surfaces, (B) brushes grafted from gold coated silicon surfaces and (C) brushes grafted from gold colloids on atomic force microscopy probes. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/52285/52285fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/52285/52285fig3highres.jpg" /> 
Figure 3. FTIR spectra of the PMMA (blue) and PNIPAM (green) brushes on silicon (thick lines) and gold (thin lines). The data was taken from Suppl. Mat. of Ref. 22. PMMA wavenumbers (cm&#8722;1): 3,050&#8211;2,990 (CH stretching vibration), 1,730 C=O (double bond stretching vibration), 1,450 (CH3 and CH2 deformation vibration), 1,260&#8211;1,040 (C-O-C single bond stretching vibration), 880&#8211;960 (C-O-C single bond deformation vibration). At 1,730 cm&#8722;1 the characteristic stretching vibration peak of the C=O group is apparent. PNIPAM wavenumbers (cm-1): 3,289 (N-H symmetric and asymmetric stretching vibration), 3,078, 2,971, 2,933, 2,874 (asymmetric and symmetric C-H stretching vibration in -CH2-), 1,635 (C=O stretching vibration), 1,535 (amide II), 1,458 (C-H asymmetric bending deformations), 1,386 (C-H symmetric bending deformations), 1,366&#8211;1,170 (C-N asymmetric stretching vibrations). At 1,635 and 1,535 cm&#8722;1 the characteristic stretching vibration peaks of the amide group are apparent.
<img alt="Figure 4" src="/files/ftp_upload/52285/52285fig4highres.jpg" width="700px" /> 
Figure 4. Averaged, filtered and smoothed force traces upon sliding the miscible (left) and immiscible (right) systems (adjusted from Ref. 22). The surface is moved back and forth by 40 &#956;m at a scan-rate of 1 Hz and normal load of 30 nN.

Watch this Scientific Journal Video about Preparation and Friction Force Microscopy Measurements of Immiscible, Opposing Polymer Brushes at JoVE.com

Preparation and Friction Force Microscopy Measurements of Immiscible, Opposing Polymer Brushes

The methodology to perform friction force microscopy experiments for contacting brushes is presented: Two polymer brushes that are grafted from (a) substrates and (b) colloidal probes are slid to show that, by using two contacting immiscible brush systems, friction in sliding contacts is reduced compared to miscible brush systems.

Solvated polymer brushes are well known to lubricate high-pressure contacts, because they can sustain a positive normal load while maintaining low ...

preparation-friction-force-microscopy-measurements-immiscible

Universidad de Cartagena UDEC

Research

JoVE Journal

Engineering

14.0K Views.  Forschungszentrum Jülich. The methodology to perform friction force microscopy experiments for contacting brushes is presented: Two polymer brushes that are grafted from (a) substrates and (b) colloidal probes are slid to show that, by using two contacting immiscible brush systems, friction in sliding contacts is reduced compared to miscible brush systems.

Video: Preparation and Friction Force Microscopy Measurements of Immiscible, Opposing Polymer Brushes

Confocal Imaging of Confined Quiescent and Flowing Colloid-polymer Mixtures

The behavior of confined colloidal suspensions with attractive interparticle interactions is critical to the rational design of materials for directed assembly1-3, drug delivery4, improved hydrocarbon recovery5-7, and flowable electrodes for energy storage8. Suspensions containing fluorescent colloids and non-adsorbing polymers are appealing model systems, as the ratio of the polymer radius of gyration to the particle radius&#160;and concentration of polymer control the range and strength of the interparticle attraction, respectively. By tuning the polymer properties and the volume fraction of the colloids, colloid fluids, fluids of clusters, gels, crystals, and glasses can be obtained9.&#160;Confocal microscopy, a variant of fluorescence microscopy, allows an optically transparent and fluorescent sample to be imaged with high spatial and temporal resolution in three dimensions. In this technique, a small pinhole or slit blocks the emitted fluorescent light from regions of the sample that are outside the focal volume of the microscope optical system. As a result, only a thin section of the sample in the focal plane is imaged. This technique is particularly well suited to probe the structure and dynamics in dense colloidal suspensions at the single-particle scale: the particles are large enough to be resolved using visible light and diffuse slowly enough to be captured at typical scan speeds of commercial confocal systems10. Improvements in scan speeds and analysis algorithms have also enabled quantitative confocal imaging of flowing suspensions11-16,37.&#160;In this paper, we demonstrate confocal microscopy experiments to probe the confined phase behavior and flow properties of colloid-polymer mixtures. We first prepare colloid-polymer mixtures that are density- and refractive-index matched. Next, we report a standard protocol for imaging quiescent dense colloid-polymer mixtures under varying confinement in thin wedge-shaped cells. Finally, we demonstrate a protocol for imaging colloid-polymer mixtures during microchannel flow.

Confocal microscopy is used to image quiescent and flowing colloid-polymer mixtures, which are studied as model systems for attractive suspensions. Image analysis algorithms are used to calculate structural and dynamic metrics for the colloidal particles that measure changes due to geometric confinement.

The behavior of confined colloidal suspensions with attractive interparticle interactions is critical to the rational design of materials for directed ...

Chemistry

Quantitative and Qualitative Examination of Particle-particle Interactions Using Colloidal Probe Nanoscopy

Colloidal Probe Nanoscopy (CPN), the study of the nano-scale interactive forces between a specifically prepared colloidal probe and any chosen substrate using the Atomic Force Microscope (AFM), can provide key insights into physical interactions present within colloidal systems. Colloidal systems are widely existent in several applications including, pharmaceuticals, foods, paints, paper, soil and minerals, detergents, printing and much more.1-3 Furthermore, colloids can exist in many states such as emulsions, foams and suspensions. Using colloidal probe nanoscopy one can obtain key information on the adhesive properties, binding energies and even gain insight into the physical stability and coagulation kinetics of the colloids present within. Additionally, colloidal probe nanoscopy can be used with biological cells to aid in drug discovery and formulation development. In this paper we describe a method for conducting colloidal probe nanoscopy, discuss key factors that are important to consider during the measurement, and show that both quantitative and qualitative data that can be obtained from such measurements.

Colloidal probe nanoscopy can be used within a variety of fields to gain insight into the physical stability and coagulation kinetics of colloidal systems and aid in drug discovery and formulation sciences using biological systems. The method described within provides a quantitative and qualitative means to study such systems.

Colloidal Probe Nanoscopy (CPN), the study of the nano-scale interactive forces between a specifically prepared colloidal probe and any chosen substrate ...

Multiplexed Single-molecule Force Proteolysis Measurements Using Magnetic Tweezers

The generation and detection of mechanical forces is a ubiquitous aspect of cell physiology, with direct relevance to cancer metastasis1, atherogenesis2 and wound healing3. In each of these examples, cells both exert force on their surroundings and simultaneously enzymatically remodel the extracellular matrix (ECM). The effect of forces on ECM has thus become an area of considerable interest due to its likely biological and medical importance4-7.
	Single molecule techniques such as optical trapping8, atomic force microscopy9, and magnetic tweezers10,11 allow researchers to probe the function of enzymes at a molecular level by exerting forces on individual proteins. Of these techniques, magnetic tweezers (MT) are notable for their low cost and high throughput. MT exert forces in the range of ~1-100 pN and can provide millisecond temporal resolution, qualities that are well matched to the study of enzyme mechanism at the single-molecule level12. Here we report a highly parallelizable MT assay to study the effect of force on the proteolysis of single protein molecules. We present the specific example of the proteolysis of a trimeric collagen peptide by matrix metalloproteinase 1 (MMP-1); however, this assay can be easily adapted to study other substrates and proteases.

In this article we describe the use of magnetic tweezers to study the effect of force on enzymatic proteolysis at the single molecule level in a highly parallelizable manner.

The  generation and detection of mechanical forces is a ubiquitous aspect of cell physiology,  with direct relevance to cancer metastasis1, atherogenesis2 ...

Bioengineering

Investigating Single Molecule Adhesion by Atomic Force Spectroscopy

Atomic force spectroscopy is an ideal tool to study molecules at surfaces and interfaces. An experimental protocol to couple a large variety of single molecules covalently onto an AFM tip is presented. At the same time the AFM tip is passivated to prevent unspecific interactions between the tip and the substrate, which is a prerequisite to study single molecules attached to the AFM tip. Analyses to determine the adhesion force, the adhesion length, and the free energy of these molecules on solid surfaces and bio-interfaces are shortly presented and external references for further reading are provided. Example molecules are the poly(amino acid) polytyrosine, the graft polymer PI-g-PS and the phospholipid POPE (1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine). These molecules are desorbed from different surfaces like CH3-SAMs, hydrogen terminated diamond and supported lipid bilayers under various solvent conditions. Finally, the advantages of force spectroscopic single molecule experiments are discussed including means to decide if truly a single molecule has been studied in the experiment.

A protocol to couple a large variety of single molecules covalently onto an AFM tip is presented. Procedures and examples to determine the adhesion force and free energy of these molecules on solid supports and bio-interfaces are provided.

Atomic force spectroscopy is an ideal tool to study molecules at surfaces and interfaces. An experimental protocol to couple a large variety of single ...

Prescribed 3-D Direct Writing of Suspended Micron/Sub-micron Scale Fiber Structures via a Robotic Dispensing System

A 3-axis dispensing system is utilized to control the initiating and terminating fiber positions and trajectory via the dispensing software. The polymer fiber length and orientation is defined by the spatial positioning of the dispensing system 3-axis stages. The fiber diameter is defined by the prescribed dispense time of the dispensing system valve, the feed rate (the speed at which the stage traverses from an initiating to a terminating position), the gauge diameter of the dispensing tip, the viscosity and surface tension of the polymer solution, and the programmed drawing length. The stage feed rate affects the polymer solution&#8217;s evaporation rate and capillary breakup of the filaments. The dispensing system consists of a pneumatic valve controller, a droplet-dispensing valve and a dispensing tip. Characterization of the direct write process to determine the optimum combination of factors leads to repeatedly acquiring the desired range of fiber diameters. The advantage of this robotic dispensing system is the ease of obtaining a precise range of micron/sub-micron fibers onto a desired, programmed location via automated process control. Here, the discussed self-assembled micron/sub-micron scale 3D structures have been employed to fabricate suspended structures to create micron/sub-micron fluidic devices and bioengineered scaffolds.

Here, we present a protocol to fabricate freely-suspended, micron/sub-micron scale polymer fibers and &#8220;web-like&#8221; structures generated via automated direct writing procedure by means of a 3-axis dispensing system.

A 3-axis dispensing system is utilized to control the initiating and terminating fiber positions and trajectory via the dispensing software. The polymer ...

Self-assembling Morphologies Obtained from Helical Polycarbodiimide Copolymers and Their Triazole Derivatives

A facile method for the preparation of polycarbodiimide-based secondary structures (e.g., nano-rings, &#34;craters,&#34; fibers, looped fibers, fibrous networks, ribbons, worm-like aggregates, toroidal structures, and spherical particles) is described. These aggregates are morphologically influenced by extensive hydrophobic side chain-side chain interactions of the singular polycarbodiimide strands, as inferred by atomic force microscopy (AFM) and scanning electron microscopy (SEM) techniques. Polycarbodiimide-g-polystyrene copolymers (PS-PCDs) were prepared by a combination of synthetic methods, including coordination-insertion polymerization, copper(I)-catalyzed azide alkyne cycloaddition (CuAAC) &#34;click&#34; chemistry, and atom transfer radical polymerization (ATRP). PS-PCDs were found to form specific toroidal architectures at low concentrations in CHCl3. To determine the influence of a more polar solvent medium (i.e., THF and THF/EtOH) on polymer aggregation behavior, a number of representative PS-PCD composites have been tested to show discrete concentration-dependent spherical particles. These fundamental studies are of practical interest to the development of experimental procedures for desirable architectures by directed self-assembly in thin film. These architectures may be exploited as drug carriers, whereas other morphological findings represent certain interest in the area of novel functional materials.

Here, we present a protocol to prepare and visualize secondary structures (e.g., fibers, toroidal architectures, and nano-spheres) derived from helical polycarbodiimides. The morphology characterized by both atomic force microscopy (AFM) and scanning electron microscopy (SEM) was shown to depend on molecular structure, concentration, and the solvent of choice.

A facile method for the preparation of polycarbodiimide-based secondary structures (e.g., nano-rings, &#34;craters,&#34; fibers, looped fibers, fibrous ...

Sample Preparation in Quartz Crystal Microbalance Measurements of Protein Adsorption and Polymer Mechanics

In this study, we present various examples of how thin film preparation for quartz crystal microbalance experiments informs the appropriate modeling of the data and determines which properties of the film can be quantified. The quartz crystal microbalance offers a uniquely sensitive platform for measuring fine changes in mass and/or mechanical properties of an applied film by observing the changes in mechanical resonance of a quartz crystal oscillating at high frequency. The advantages of this approach include its experimental versatility, ability to study changes in properties over a wide range of experimental time lengths, and the use of small sample sizes. We demonstrate that, based on the thickness and shear modulus of the layer deposited on the sensor, we can acquire different information from the material. Here, this concept is specifically exploited to display experimental parameters resulting in mass and viscoelastic calculations of adsorbed collagen on gold and polyelectrolyte complexes during swelling as a function of salt concentration.

The quartz crystal microbalance can provide accurate mass and viscoelastic properties for films in the micron or submicron range, which is relevant for investigations in biomedical and environmental sensing, coatings, and polymer science. The sample thickness influences which information can be obtained from the material in contact with the sensor.

In this study, we present various examples of how thin film preparation for quartz crystal microbalance experiments informs the appropriate modeling of ...

Covalent Attachment of Single Molecules for AFM-based Force Spectroscopy

Atomic force microscopy (AFM)-based single molecule force spectroscopy is an ideal tool for investigating the interactions between a single polymer and surfaces. For a true single molecule experiment, covalent attachment of the probe molecule is essential because only then can hundreds of force-extension traces with one and the same single molecule be obtained. Many traces are in turn necessary to prove that a single molecule alone is probed. Additionally, passivation is crucial for preventing unwanted interactions between the single probe molecule and the AFM cantilever tip as well as between the AFM cantilever tip and the underlying surface. The functionalization protocol presented here is reliable and can easily be applied to a variety of polymers. Characteristic single molecule events (i.e., stretches and plateaus) are detected in the force-extension traces. From these events, physical parameters such as stretching force, desorption force and desorption length can be obtained. This is particularly important for the precise investigation of stimuli-responsive systems at the single molecule level. As exemplary systems poly(ethylene glycol) (PEG), poly(N-isopropylacrylamide) (PNiPAM) and polystyrene (PS) are stretched and desorbed from SiOx (for PEG and PNiPAM) and from hydrophobic self-assembled monolayer surfaces (for PS) in aqueous environment.

Covalent attachment of probe molecules to atomic force microscopy (AFM) cantilever tips is an essential technique for the investigation of their physical properties. This allows us to determine the stretching force, desorption force and length of polymers via AFM-based single molecule force spectroscopy with high reproducibility.

Atomic force microscopy (AFM)-based single molecule force spectroscopy is an ideal tool for investigating the interactions between a single polymer and ...

Atomic Force Microscopy Cantilever-Based Nanoindentation: Mechanical Property Measurements at the Nanoscale in Air and Fluid

An atomic force microscope (AFM) fundamentally measures the interaction between a nanoscale AFM probe tip and the sample surface. If the force applied by the probe tip and its contact area with the sample can be quantified, it is possible to determine the nanoscale mechanical properties (e.g., elastic or Young&#39;s modulus) of the surface being probed. A detailed procedure for performing quantitative AFM cantilever-based nanoindentation experiments is provided here, with representative examples of how the technique can be applied to determine the elastic moduli of a wide variety of sample types, ranging from kPa to GPa. These include live mesenchymal stem cells (MSCs) and nuclei in physiological buffer, resin-embedded dehydrated loblolly pine cross-sections, and Bakken shales of varying composition.
Additionally, AFM cantilever-based nanoindentation is used to probe the rupture strength (i.e., breakthrough force) of phospholipid bilayers. Important practical considerations such as method choice and development, probe selection and calibration, region of interest identification, sample heterogeneity, feature size and aspect ratio, tip wear, surface roughness, and data analysis and measurement statistics are discussed to aid proper implementation of the technique. Finally, co-localization of AFM-derived nanomechanical maps with electron microscopy techniques that provide additional information regarding elemental composition is demonstrated.

Quantifying the contact area and force applied by an atomic force microscope (AFM) probe tip to a sample surface enables nanoscale mechanical property determination. Best practices to implement AFM cantilever-based nanoindentation in air or fluid on soft and hard samples to measure elastic modulus or other nanomechanical properties are discussed.

An atomic force microscope (AFM) fundamentally measures the interaction between a nanoscale AFM probe tip and the sample surface. If the force applied by ...

Protrusion Force Microscopy: A Method to Quantify Forces Developed by Cell Protrusions

In numerous biological contexts, animal cells need to interact physically with their environment by developing mechanical forces. Among these, traction forces have been well-characterized, but there is a lack of techniques allowing the measurement of the protrusion forces exerted by cells orthogonally to their substrate. We designed an experimental setup to measure the protrusion forces exerted by adherent cells on their substrate. Cells plated on a compliant Formvar sheet deform this substrate and the resulting topography is mapped by atomic force microscopy (AFM) at the nanometer scale. Force values are then extracted from an analysis of the deformation profile based on the geometry of the protrusive cellular structures. Hence, the forces exerted by the individual protruding units of a living cell can be measured over time. This technique will enable the study of force generation and its regulation in the many cellular processes involving protrusion. Here, we describe its application to measure the protrusive forces generated by podosomes formed by human macrophages.

Here, we detail the experimental techniques used to evaluate the protrusion forces that podosomes apply on a compliant film, from the preparation of the film to the automated analysis of topographical images.

In numerous biological contexts, animal cells need to interact physically with their environment by developing mechanical forces. Among these, traction ...