The authors thank Swiss National Foundation for Science (SNF) for the financial support (grant number P2ELP2_162116 and P300P2_171219), the Darwin College, Erasmus+ program for the financial support (grant number 2018-1-LT01-KA103-046719-15400-P3) and the research leading to these results has received funding from the European Research Council under the European Union&#39;s Seventh Framework Programme (FP7/2007-2013) through the ERC grant PhysProt (agreement number 337969), the Newman Foundation (T.P.J.K.) and The Cambridge Centre for Misfolding Diseases (C.G., M.V., and T.P.J.K.).

1. Aggregation assays on fluorescence plate readers
NOTE: The protocol described here is an example of how to study the aggregation of any protein or peptide by chemical kinetics. In particular, it describes an optimized protocol to study the aggregation of the A&#946;42 peptide, which is involved in the onset and progression of Alzheimer&#8217;s disease58,59. A similar protocol can be adjusted and adopted towards studying the aggregation of any protein or peptide.
<ol>
	<li>Obtain a highly pure monomeric solution of A&#946;42 through ion-exchange and size-exclusion chromatography techniques to distinguish protein fractions containing A&#946;42, and to isolate the monomeric fraction from other aggregated forms of A&#946;42 obtained58,59.</li>
	<li>Dilute the A&#946;42 peptide in 20 mM sodium phosphate buffer, 200 &#181;M EDTA at pH 8.058 to a desired final concentration ranging between 1&#8722;5 &#181;M, in 1.5 mL low-bind tubes. Samples collected for AFM measurements should not contain the fluorescent dye used in the aggregation assay (thioflavin T, ThT), as it might introduce artefacts during sample deposition for AFM analysis. Thus, prepare triplicates of two identical solutions: i) the first containing monomeric A&#946;42 to follow the process of aggregation by AFM; ii) the second containing the A&#946;42 monomeric with addition of 20 &#181;M ThT as a tracer to monitor the kinetics of aggregation. 
	NOTE: It is important to perform steps 1.1 and 1.2 on ice. This procedure is to ensure that the monomeric A&#946;42 solution does not aggregate until initiated in the plate reader. When handling samples, careful pipetting is essential to avoid the introduction of any air bubbles that may affect the protein sample.</li>
	<li>Pipette 80 &#181;L of each sample in each well of a 96-well, half-area plate of black polystyrene with clear bottom and nonbinding surfaces.</li>
	<li>Seal the plate with a foil to minimize the evaporation of the sample over the course of the aggregation.</li>
	<li>Equilibrate the temperature of the plate reader to 37 &#176;C.</li>
	<li>Set up the aggregation protocol to read fluorescence measurements at fixed time-point intervals, at an excitation wavelength of 440 nm and an emission wavelength of 480 nm. The aggregation should be initiated in quiescent conditions.</li>
	<li>Insert the plate into the plate reader. 
	NOTE: It is important to be careful while handling the plate, to ensure that the aggregation is not triggered prior to starting the plate reader. Use the gain adjustment settings to ensure the optimal readout of the fluorescence of each sample.</li>
	<li>Start the measurement. 
	NOTE: The aggregation experiment is concluded when the ThT fluorescence sigmoidal curve reaches a plateau58.</li>
</ol>
2. Sample preparation for AFM and nano-IR measurements
<ol>
	<li>Glue highest quality grade V1 mica disc to high quality magnetic stainless-steel disc using adhesive tabs or double-sided tape.</li>
	<li>Etch mica by placing a piece of adhesive tape onto the mica surface and pulling off the top layer of the mica. This procedure will produce clean, atomically flat surface, suitable for sample deposition. 
	NOTE: Etched surface must be uniform and smooth. 
	CAUTION: Do not touch or breathe directly above freshly etched mica, as this will introduce artefacts.</li>
	<li>Pause/stop the plate reader measurement to collect the time point of interest during the aggregation process of the protein.</li>
	<li>Remove the sealing foil and collect 10 &#181;L aliquot of the samples without ThT from the well into a 1.5 mL low-bind tubes.</li>
	<li>Deposit the 10 &#181;L of the sample on the freshly etched mica. For AFM-IR measurements samples must be deposited on freshly stripped gold substrate.</li>
	<li>Incubate the solution on the mica for 1 min to allow physisorbtion. 
	NOTE: Longer incubation time would allow better absorption on the surface but may induce artificial self-organization and self-assembly25. To avoid these effects, spray deposition may be exploited25.</li>
	<li>Rinse three times with 1 mL of ultrapure water.</li>
	<li>Dry under a gentle flow of nitrogen to measure the sample in air environment.</li>
</ol>
3. AFM imaging of the morphology of protein aggregates
NOTE: Morphology measurements can be performed both in contact and dynamic mode, in the following steps the latter is described since it reduces lateral forces to measure the 3D morphology of the sample with high resolution. AFM-IR measurements will be performed in contact mode to enhance AFM-IR signal-to-noise ratio.
<ol>
	<li>Turn on the AFM system at least 30 min before the measurements in order to enable the system to reach thermal stability. Click on Setup, then on Probe. In Probe Change window click on Next to prepare Z, Focus Stage.</li>
	<li>Turn off the AFM beam switch (if it is on). Unlock the dovetail locks, disconnect the head from the system and in Probe Change window click on Next.</li>
	<li>Mount the AFM cantilever on the probe holder. Connect the head to the system and lock the dovetail locks. 
	NOTE: Optimal cantilevers to study biological samples in dynamic mode AFM have spring constant ranging between 2&#8722;40 N m-1 and an apex radii ranging between 2&#8722;8 nm14.</li>
	<li>Turn on the AFM laser beam switch and in Probe Change window click on Next.</li>
	<li>In the pop-up window click on No if the cantilever type did not change. Adjust optical alignment knobs to find the cantilever. Adjust focus onto cantilever and click on Next.</li>
	<li>In the pop-up window click on Yes if the focus is on the cantilever.</li>
	<li>Position the laser beam at the end of the cantilever using the knobs controlling position of the detection laser. Maximize the total signal measured by the four-quadrant photodiode to at least &#62;1 V using the knobs controlling the position of the deflected laser beam on the position sensitive photodiode (PSPD). In Probe Change window click on Next and then on Close.</li>
	<li>Wait 15 min for the cantilever to reach thermal stability. Readjust the position of the deflected laser beam on the PSPD if necessary.</li>
	<li>Click on NCM SWEEP, choose desired amplitude of oscillation, click on Use Phase, click on auto and tune the cantilever close to the maximum of its first free resonance of oscillation, which is of approximately 300 kHz for a cantilever with a spring constant of 40 N m-1. 
	NOTE: Tuning the cantilever out of the maximum of its free resonance assures higher stability of the measurements14.</li>
	<li>Place the sample on the sample holder. Choose suitable imaging parameters. Typical scanning rate is 0.3&#8722;1.0 Hz for a scan area of 1 x 1 &#181;m2 to 5 x 5 &#181;m2. Typical resolution needed is between 256 x 256 and 1024 x 1024 pixels. Click on Scan Area to choose the scan size and the pixel number.</li>
	<li>Focus the optical view on the sample. Click on Approach to approach the sample surface. Once approach is completed click on Lift 100 &#181;m to rise the AFM tip 100 &#181;m above the surface of the sample. Click Expand on the optical image of the sample. Click on the Focus Stage bar to focus the view on the surface of the sample.</li>
	<li>Use the arrows to move in the region of the sample of interest. Engage the surface by pressing Approach button. Click on Line Scan button, and check if the tip is following the surface well, if necessary, adjust the Set Point.</li>
	<li>Start imaging the sample surface by pressing Scan button. During imaging, to avoid large imaging force and keep consistency of within independent samples, maintain a constant regime of phase change not exceeding &#916;20&#176;42.</li>
	<li>Enter file name and choose base directory where the acquired image will be saved.</li>
</ol>
4. Infrared nanospectroscopy measurements of protein aggregates
<ol>
	<li>Turn on the AFM-IR system and the infrared (IR) laser60 30&#8722;60 min before measurements for thermal stabilization. Typical lasers for AFM-IR instruments are optical parameter oscillators (OPO) and quantum cascade lasers (QCL).</li>
	<li>Open the built-in software to control the instrument. Click on file and then on new to open a new nanoIR file. Press the button initialise to start the AFM-IR system.</li>
	<li>Open the instrument cover and mount on the AFM-IR system a silicon gold coated probe with a nominal radius of 30 nm and a spring constant of 0.2 N/m to measure the sample in contact mode.</li>
	<li>Click on Load in the section AFM probe and then next. In focus on probe section click on the arrows to focus the camera on the cantilever. In the section sample XY movement, use the arrows to place the cross-air at the end of the cantilever.</li>
	<li>Rotate the knobs controlling the position of the detection laser to position laser at the end of the cantilever. Rotate the laser knobs to detect and maximize the sum measured by the four-quadrant photodiode to a value &#62;3 V. Rotate the deflection knob to adjust the cantilever deflection to -1 V and then click next. Close the cover of the instrument.</li>
	<li>In the section focus on sample use the arrows to focus the camera on the sample. Then, in the section sample XY movement use the arrows to move in the region of interest of the sample and click on approach and engage.</li>
	<li>In the microscope window select as inputs the channels of interest to map the biophysical properties of the sample. In particular, choose height for morphology, Amplitude 2 for IR absorption and PLL frequency to map tip-sample contact resonance.</li>
	<li>In the AFM scan section of the controls window, use similar parameters as in section 3 (i.e., scan rate 0.1&#8722;1.0 Hz, between 256 x 256 and 1024 x 1024 pixels). Choose as value of the gains according to sample roughness: integral I = 1&#8722;10 and proportional P = 10&#8722;30. Then, click on scan to acquire a morphology map. 
	NOTE: The morphology can be similarly measured in the dynamic tapping mode, but AFM-IR measurements are performed here in the contact mode to have higher signal to noise ratios.</li>
	<li>After the mapping of morphology is finished, in the microscope window, click on the height map to position the probe on the top of one aggregate. Then in the nanoIR section of the controls window, click on start IR to illuminate the sample with the IR laser.</li>
	<li>To focus the infrared laser on the cantilever, click on optimisation. In this window, write a wavenumber where the sample is going to have high absorbance and click add. For protein, a typical value is 1655 cm-1. Click on scan to find the IR laser position and click on update to align its position with the cantilever. Close the window.</li>
	<li>In the section general of the nanoIR setting, write a wavenumber where high absorbance in the relative field is expected. Then, deactivate the Band Pass Filter option and look at the meter reading and at the fast Fourier transform (FFT) of the cantilever response. In the FFT window move the green cursor to read the resonance frequency of the cantilever. A typical value of the FFT of the resonance of the cantilevers is around 300 kHz. Write this resonance frequency value in the general section in the freq. centre field and use a freq. window of 50 kHz. 
	NOTE: Select a laser power that is low enough to not saturate the meter reading and to have distortion in the cantilever response and to not overheat the sample.</li>
	<li>Click on laser pulse tune window to use the resonance enhanced mode. Choose a frequency centre of 300 kHz, a tune range of 50 kHz and a duty cycle of the laser of 5%. Click on acquire to sweep the pulse rate of the laser. By using the cursor in the graph, tune the laser pulse to the frequency of the mechanical response of the thermal expansion of the sample absorbing the IR light. Then, select the option PLL to monitor the contact resonance between the sample and the tip. Press the zero button in the PLL window and tick enable to track the sample-tip contact resonance. Choose an integral gain I = 0.5 and proportional gain P = 10. Close the window.</li>
	<li>Open again the optimisation window and find the position of the IR laser for at least 3 wavenumbers corresponding to major absorbance bands of the sample (amide band I-II-III) and for at least one wavenumber for each chip of the laser.</li>
	<li>Click on Tools | IR Background Calibration | New. In the window select the spectroscopic region of interest (1800&#8722;1200 cm-1 to study protein samples); choose the same pulse rate as determined in step 4.12 and a duty cycle of 5%; click on fast acquisition and select the laser speed, typical range for a quantum cascade laser is between 20&#8722;500 cm-1. Click on acquire to measure the IR laser background. This background spectrum will be used for normalization of measured nanoscale localized spectra. Close the window. 
	NOTE: If a fast laser and resonance enhanced mode is not available, skip step 4.12 and select stepped spectra instead of fast in both the background and IR spectra acquisition windows. However, single aggregate sensitivity will not be reached.</li>
	<li>In the IR spectra settings, choose an IR spectrum resolution between 1&#8722;4 cm-1 and a number of co-averages of at least 64x. Click on acquire to measure a nanoscale localized IR spectrum in the protein range (1800&#8722;1200 cm-1).</li>
	<li>To acquire a nanoscale resolved chemical map, select IR imaging option, choose a wavenumber of interest (e.g., 1655 cm-1 for amide band I) and click on scan in the AFM scan window.</li>
	<li>Once the mapping is completed, go to file and save the measurements. To analyze the acquired maps of morphology, contact-resonance and chemistry, as well as the nanoscale localized spectra, use the built-in AFM image processing software. Spectra and Images can be saved as .csv or .axz files for further detailed analysis with commercial software.</li>
</ol>
5. Image processing and analysis of cross-sectional dimensions
<ol>
	<li>Flatten raw images14,17,19,41,42,59 using built-in AFM image processing software or commercial software. 
	NOTE: The aggregates should be masked from the calculation to avoid artefacts of analysis and underestimation of their measured height.</li>
	<li>Flatten the image by 0 order plane fit subtraction.</li>
	<li>Flatten the image by plane and then line by line at a 1st regression order, the second step is repeated until the flat baseline in line profile of the image is reached.</li>
	<li>If the sample is very crowded, contains exceptionally high aggregates or scanner bow artefact is present, flatten image using 2nd regression order fit.</li>
	<li>Measure aggregate height and width from a line profile taken perpendicular to the fibril axis14,17,19,41,42,59.</li>
	<li>Measure fibril length parallel to the fibril axis14,17,19,41,42,59.</li>
</ol>

The authors have nothing to disclose.

The first critical step in this protocol is the preparation of monomeric proteins, such as in the case of A&#946;42 solution described in steps 1.1 and 1.2. It is essential to initiate the aggregation process from a highly pure, monomeric solution, as the presence of oligomeric or aggregated species may result in poor reproducibility of the aggregation kinetics58, and induce artefacts in the AFM measurements (e.g., fibrillar species will be evident at the initial stages of the aggregation), which may lead to the misinterpretation of the data. Highly reproducible kinetics data of amyloid formation based on ThT fluorescence assay, in association with the master equation formalism of chemical kinetics5,6,7, have allowed to define the A&#946;42 aggregation mechanism in terms of its underlying molecular events. Chemical kinetics connects the microscopic steps underlying amyloid formation with their macroscopic manifestations by considering the different ways in which new aggregates can form and grow, which are for instance elongation at the aggregate ends or secondary nucleation on the aggregate surface. However, chemical kinetics by itself does not directly enable the visualization of the possible nucleation phenomena at the nanoscale requiring their combination with single molecule methods.
The second critical step in this protocol is the substrate preparation and sample deposition procedures described in steps 2.2 and 2.6&#8722;2.8. To avoid artefacts, the sample must be deposited on a clean, atomically flat surface. Proper etching of the mica is essential to achieve artefact-free, high-resolution in AFM measurements. The sample deposition time is also extremely important, as longer incubation time allows better absorption on the substrate surface. However, it might also induce artificial self-organization and self-assembly25, which may induce artefacts (e.g., surface-induced aggregate species) that may lead to data misinterpretation. In addition, the mica surface is negatively charged, which means that only positively charged molecules easily absorb on it. If the net charge of the sample is negative, the surface of the mica can be positively functionalized using APTES for a better phisisorption23. Microfluidic spray deposition25 can be exploited to avoid these effects and artefacts and deposit the sample in a single step and artefact-free manner.
The third critical step is the proper setup and the choice of imaging parameters for sample imaging via AFM and AFM-IR described in section 3. The AFM tips used for the sample imaging should be sharp enough (apex radii of 2&#8722;8 nm) to achieve high-resolution and minimize convolution effects (broadening of the sample features by the tip)14, which may induce uncertainties in the image of the sample. The choice of imaging mode, contact or dynamic, is also important. For sample imaging via conventional AFM, the dynamic mode is preferred over the contact mode as the latter mode induces large lateral tip-sample frictional forces that may cause sample damage and introduce artefacts in the measurements (e.g., reduction in the sample height due to nanoindentation)14,61,62. Conversely, the contact mode is preferred for the measurements via nano-IR to enhance AFM-IR signal. For measurements in dynamic mode, the cantilever should be tuned just slightly below (tapping mode) or above (non-contact mode) the maximum of its first free resonance of oscillation to assure higher stability of the measurements14. The imaging resolution, which depends on the pixel number and scan area, should be high enough to capture the smallest degree of detail present in a specimen (e.g., 1024 x 1024 pixels for a 4 x 4 &#181;m2 area)14,63. Low imaging resolution can induce distortions and uncertainties in the image of the sample due to the loss of the vertical and lateral information upon digitisation of the signal14. The scan rate, used for imaging, should be low enough for the tip to be able to follow surface features properly as well as to have enough time to acquire chemical information14. One of the most important imaging parameters is the imaging force. In contact mode, it is fundamental to use a low interaction force to preserve structure of the sample. In dynamic mode, the energy dissipation on the samples should be kept constant in order to consistently compare morphology of distinct samples; consistent imaging of independent samples can be obtained by maintaining a constant regime of a phase change not exceeding &#916;20&#176;14,42. Large imaging forces should be avoided as they may induce distortions and uncertainties in the sample images.
Thermal drift caused by the expansion and contraction of AFM parts due to thermal fluctuations can induce distortions and artefacts in the image of the sample64,65. Drifts in the vertical direction may cause the cantilever to lose track of the surface as well as to crash into the surface, while drifts in the lateral direction usually result in elongation of the surface features and image distortion, which makes it difficult to achieve precise measurements of the sample features. The effect of these thermal drifts can be minimized by accurate temperature control of the laboratory as well as giving enough time (around 30 min) for the system to become stable or by performing fast scans66,67,68,69,70.
The quality of the nano-IR imaging and spectra collection by AFM-IR can be affected by several factors, of which the most important are: (i) a wrong division of the IR signal on the sample by the IR background, (ii) a large variation of nanomechanical contact between tip and the sample, and (iii) an excessive heating of the sample causing its softening. To correctly divide the IR spectrum of the sample by the collected background, it is crucial that they are collected at the same laser power. Indeed, the spectral line shape of the IR laser background depends on the power of the laser. Then, in order to avoid the influence of the mechanical properties of the sample into the measured chemical information, it is crucial to monitor and track the contact resonance between the sample and the tip during spectra and image acquisition. For spectra acquisition, ideally, it is sufficient to pulse the laser at a fixed contact resonance. However, if the spectrum is acquired on a large spectroscopic range, high intensity peaks could cause strong heating of the sample and its softening, thus changing the tip-sample contact resonance and artificially reducing the IR peak amplitude. For this reason, it is important to track contact resonance during spectra acquisition in order to verify that the spectrum is not affected by excessive heating of the sample.
In conclusion, conventional AFM and nano-IR are capable of investigating with high resolution the morphological, structural and chemical properties of the individual species forming during protein aggregation24,38. However, they lack the capability of chemical kinetics to follow their rapid kinetics of formation in native bulk conditions. In order to unravel the conformational changes that protein undergo during their aggregation and misfolding, it is necessary to develop and apply novel biophysical methods capable of bringing together the capabilities of bulk biophysical methods with the investigation of the heterogeneity and ultrastructural properties of protein aggregation at the nanoscale. This approach represents a fruitful avenue to address the challenge of understanding the problem of protein self-assembly and its role in health and disease. Indeed, single aggregate approaches are capable to unravel and elucidate the molecular mechanisms of protein aggregation polymorphism and formation. This information is central to address the challenge of understanding protein aggregation and its role in the onset and progression of human diseases, as well as understanding their biophysical properties for biotechnology applications.

Over 40 million people worldwide are currently affected by neurodegenerative disorders, such as Alzheimer&#8217;s (AD)1 and Parkinson&#8217;s (PD)2 diseases. More generally, more than fifty pathologies are associated at the molecular level with protein misfolding and aggregation, a process that leads to the proliferation of insoluble fibrillar protein aggregates, known as amyloid deposits3,4. The molecular origins of neurodegeneration and its links with protein conformational changes of proteins leading to amyloid formation, however, remain unclear, in large part because of the high level of heterogeneity, transient nature and nanoscale dimensions of the pathological aggregates4,5.
Highly successful investigations of protein structures in the last several decades have been based widely on the use of bulk methods, including X-ray crystallography, cryo-electron microscopy and nuclear magnetic resonance spectroscopy5,6,7,8,9. Within this class of techniques, infrared (IR) spectroscopy has emerged as a sensitive analytical tool to unravel the chemical properties of biological systems such as proteins8. IR methods allow the quantification of protein secondary and quaternary structural changes during their misfolding and aggregation. In addition, in order to further decipher at the microscopic level the mechanistic details involved in the complex free energy landscapes of protein during their aggregation, a major advance has been the development of chemical kinetics tools to extend to complex self-assembly pathways including amyloid fibrils formation5,6,7,10,11,12. However, bulk spectroscopic methods provide only average information on the heterogeneous ensemble of species present in solution or involved in specific microscopic steps, thus rendering the investigation of the biophysical properties of individual aggregated species challenging at the nanoscale level13,14.
Several microscopy techniques with the capability of operating on scales smaller than the diffraction limit of light have emerged in the last decades. This class of methods includes electron microscopy (EM) and atomic force microscopy (AFM). While scanning electron microscopy (SEM) and transmission electron microscopy (TEM) provide two-dimensional (2D) images of a specimen, AFM has emerged in the last decades as a powerful and versatile technique to study three-dimensional (3D) morphologies, as well as the nanomechanical properties of a sample with sub-nanometer resolution13,14,15,16,17,18,19,20,21,22,23,24,25,26,27. The rationale behind studying protein aggregation via AFM is that this approach enables the investigation of the morphology of individual species present in solution13,14,16,17,19,20,21,25,27,28,29,30,31,32,33,34,35,36,37. In particular, by monitoring the sample as a function of time, AFM allows the investigation of the evolution of the morphology of the species within the sample, which makes it possible to follow and visualize the pathways of amyloid formation23,25,38,39,40,41,42. Furthermore, AFM enables the quantification of structural parameters such as cross-sectional heights and lengths of the individual species present in solution13,19,30,31,32,33,34,35,36,37,40,43,44,45,46,47,48. However, the study of a single biophysical property, such as morphology, is often not sufficient when studying heterogeneous and complex biological systems. AFM, SEM or TEM imaging methods alone do not readily reveal the chemical properties of heterogeneous species of amyloid aggregates at the nanoscale.
A major advance for the analysis of heterogeneous biological samples at this scale has been made recently with the development and application to the field of protein aggregation of infrared nanospectroscopy (AFM-IR)24,26,38,42,49,50,51,52. This innovative method exploits the combination of the spatial resolution of AFM (~1&#8722;10 nm) with the chemical analysis power of IR. The AFM-IR technique is based on the measurement of the photothermal induced resonance effect driven by an IR laser, and on the measurement of the thermal expansion of the sample under investigation by the AFM tip. The sample can be illuminated by the IR laser directly from the top or from the bottom in total internal reflection, similarly as in conventional infrared spectroscopy24,42,52,53. The IR laser can be pulsed with typical frequencies in the order of hundreds of kilohertz (1&#8722;1000 kHz) and tuned over a wide spectral range, typically between 1000&#8722;3300 cm-1. Although the laser source covers an area of ~30 &#181;m diameter, the spatial resolution of the AFM-IR technique is determined nominally by the AFM tip diameter, which detects the local thermal expansion of the system. AFM-IR is well suited to study biological samples because the IR signal is proportional to their thickness up to 1&#8722;1.5 &#181;m, and the resulting IR spectra are generally in agreement with the corresponding FTIR transmission spectra13,54,55. For this reason, established methods of analysis in spectroscopy can be readily applied, such as the study of chemical shifts, band shape change and de-convolution by second derivatives analysis52. Overall, combining the spatial resolution of AFM with the chemical recognition power of IR spectroscopy, AFM-IR enables the simultaneous acquisition of a wide range of morphological, mechanical and chemical properties of a sample at the nanoscale.
Here, we illustrate a protocol for the characterization of the process of protein aggregation that exploits the combination of in vitro fluorescence assays, high-resolution AFM imaging and nanoscale AFM-IR. This combined approach has already excelled in providing detailed results in studying the chemical and structural properties of individual micro-droplets formed by protein aggregates, in the study of liquid-liquid protein phase separation, and in investigating the heterogeneity and biophysical properties of individual aggregated species at the nanoscale23,26,38,45,50,53,56,57.

<table>
	<tbody>
		<tr>
			<td>AFM-IR system</td>
			<td>Anasys Instruments</td>
			<td>nanoIR 2 or 3</td>
			<td>Systems to measure thermal expansion in contact and resonance mode</td>
		</tr>
		<tr>
			<td>Corning 96-well Half Area Black/Clear Bottom Polystyrene NBS Microplate</td>
			<td>Corning</td>
			<td>3881</td>
			<td></td>
		</tr>
		<tr>
			<td>Corning Microplate Aluminium Sealing Tape</td>
			<td>Corning</td>
			<td>6570</td>
			<td></td>
		</tr>
		<tr>
			<td>Double Sided Adhesive Discs</td>
			<td>AGAR Scientific</td>
			<td>AGG3347N</td>
			<td></td>
		</tr>
		<tr>
			<td>FLUOstar Omega</td>
			<td>BMG Labtech</td>
			<td>415-101</td>
			<td>Platereader</td>
		</tr>
		<tr>
			<td>Mica Disc 10mm V1</td>
			<td>AGAR Scientific</td>
			<td>AGF7013</td>
			<td></td>
		</tr>
		<tr>
			<td>Park NX10 AFM system</td>
			<td>Park Systems</td>
			<td>N/A</td>
			<td>Atomic Force Microscope</td>
		</tr>
		<tr>
			<td>Platypus Ultra-Flat Gold Chips</td>
			<td>Platypus Technologies</td>
			<td>AU.1000.SWTSG</td>
			<td></td>
		</tr>
		<tr>
			<td>PPP-NCHR-10 cantilevers</td>
			<td>Park Systems</td>
			<td>PPP-NCHR-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Protein LowBind Tubes, 2.0mL</td>
			<td>Eppendorf</td>
			<td>30108132</td>
			<td></td>
		</tr>
		<tr>
			<td>Silicon gold coated cantilevers</td>
			<td>Anasys Instruments</td>
			<td>PR-EX-nIR2</td>
			<td></td>
		</tr>
		<tr>
			<td>SPM Specimen Discs 12mm</td>
			<td>AGAR Scientific</td>
			<td>AGF7001</td>
			<td></td>
		</tr>
	</tbody>
</table>

characterizing individual protein aggregates by infrared nanospectroscopy and atomic force microscopy

The phenomenon of protein misfolding and aggregation results in the formation of highly heterogeneous protein aggregates, which are associated with neurodegenerative conditions such as Alzheimer&#8217;s and Parkinson&#8217;s diseases. In particular low molecular weight aggregates, amyloid oligomers, have been shown to possess generic cytotoxic properties and are implicated as neurotoxins in many forms of dementia. We illustrate the use of methods based on atomic force microscopy (AFM) to address the challenging task of characterizing the morphological, structural and chemical properties of these aggregates, which are difficult to study using conventional structural methods or bulk biophysical methods because of their heterogeneity and transient nature. Scanning probe microscopy approaches are now capable of investigating the morphology of amyloid aggregates with sub-nanometer resolution. We show here that infrared (IR) nanospectroscopy (AFM-IR), which simultaneously exploits the high resolution of AFM and the chemical recognition power of IR spectroscopy, can go further and enable the characterization of the structural properties of individual protein aggregates, and thus offer insights into the aggregation mechanisms. Since the approach that we describe can be applied also to the investigations of the interactions of protein assemblies with small molecules and antibodies, it can deliver fundamental information to develop new therapeutic compounds to diagnose or treat neurodegenerative disorders.

A representative time course of A&#946;42 aggregation, as measured by the ThT fluorescence assay, is shown in Figure 1. The aggregation process is commonly characterized by a sigmoidal curve, where a lag phase is initially observed, and is followed by a steep growth phase, before the curve reaches a plateau when an equilibrium steady state is reached6,7,58. It is essential to ensure that an optimized aggregation protocol is used to generate high-quality data to study the molecular details pertaining to aggregation processes58.
High resolution of AFM enables to investigate the morphology and heterogeneity of the aggregated species at different time points of the process. During the aggregation, monitored by the ThT assay, aliquots of the sample in the plates are prepared for single aggregate investigation by AFM and nano-IR (Figure 1). The typical process flow of manual sample preparation is shown in Figure 2. At the completion of the measurement of the 3D morphology of the sample by high resolution and phase-controlled AFM, the maps are flattened to remove non-linearity of the piezoelectric scanner and reduce sources of error in post-processing analysis of sample morphology (Figure 3). Subsequently, an accurate and sensitive single molecule statistical analysis can be performed, as shown in Figure 4. From a 3D morphology map, it is possible to extract aggregate cross-sectional height, width and length (or diameter in case of spheroidal particles), which allows to distinguish and characterize distinct species of aggregates present during the aggregation time course23,38. Typical time points of interest to investigate the process of aggregation are the lag phase, the growth phase and the plateau phase (Figure 1). During the lag phase, monomeric and oligomeric species of A&#946;42 are primarily present. When visualized by AFM, monomeric and oligomeric species of A&#946;42 typically appear as spheroidal particles 1&#8722;15 nm in diameter and 0.3&#8722;2 nm in height (Figure 1, bottom left)38,39. Formation of elongated protofilaments, protofibrils and fibrils is visible during the growth phase of A&#946;42 aggregation time course (Figure 1, bottom middle)38,39. Typically, protofilaments appear as elongated features hundreds of nanometers in length and 0.5&#8722;2 nm in height, while protofibrils appear as elongated linear or curvilinear aggregates 1&#8722;5 nm in height and hundreds of nanometers in length38,39. During the plateau phase, fibrils are the dominant species of A&#946;42 aggregates. A&#946;42 fibrils typically appear as unbranched, thread-like structures, with a cross-sectional diameter of 6&#8722;10 nm and length in the order if micrometers (Figure 3, bottom right)38,39. Remarkably, this schematic representation of the morphological properties of the aggregates is a general feature of most aggregating proteins and peptides13,14.
After the investigation of the sample morphology, nano-IR can be used to investigate the chemical properties of the individual protein aggregate species present during the process of aggregation, by acquiring nanoscale-resolved IR maps and spectra both in air and native liquid environment24,26,38,4249,50,51,52,. Figure 5 shows a schematic illustration of the AFM-IR setup. An IR source is used to illuminate the sample from the bottom in total internal reflection or directly from the top as in Figure 5a. If the IR light is absorbed by the sample, it will excite the corresponding molecular vibrational energy transition levels of the species present. The vibrational energy is dissipated inside the sample in the form of thermal heating, which causes the thermal expansion of the sample. This expansion is measured at the nanoscale by the AFM tip in contact with the sample with a resolution in the order of 10 nm. At each pulse, the cantilever detects the thermal expansion. In particular, the fast expansion of the sample kicks out the cantilever from contact with the sample, and after the kick out the cantilever rings down at its natural frequencies. In order to enhance the sensitivity of AFM-IR, it is possible to tune the laser pulse frequency at the same frequency of the oscillation of the cantilever54. In order to work in this resonance-enhanced mode, it is necessary to have IR sources that can be pulsed in a wide range of frequencies, such as quantum cascade lasers that operate typically in a range between 1&#8722;1000 kHz. The peak-to-peak amplitude and the fast Fourier transform of the ringdown signal, termed IR amplitude, of the raw cantilever deflection are proportional to the IR light absorbed. These signals are detected in real time by measuring the deflection of a red laser focused on the top of the cantilever. In order to acquire chemical maps, the laser wavenumber is fixed at a certain wavenumber and the IR amplitude signal is collected at each point of the map, while, to acquire IR spectra, the position of the cantilever is fixed in a position of interest and the laser wavelength is swept along the spectroscopic range of interest. The ultimate resolution of AFM-IR enables the measurement of the chemical properties of protein aggregates with a cross-sectional height of approximately 5 nm, as represented in Figure 6.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60108/60108fig01.jpg" /> 
Figure 1: Monitoring of the aggregation time course in vitro via ThT fluorescence and AFM. 
Samples taken during the lag phase, growth phase, and plateau phase of the aggregation process as detected by ThT fluorescence were imaged via high-resolution AFM. This figure has been adapted from Ruggeri et al.38. <a href="https://www.jove.com/files/ftp_upload/60108/60108fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60108/60108fig02.jpg" /> 
Figure 2: Schematic representation of the substrate preparation and sample deposition for AFM measurements. 
(A, B) Mica etching using adhesive tape. (C) Sample deposition. (D) Sample incubation. (E) Sample rinsing with ultrapure water. (F) Sample drying under gentle flow of nitrogen. (G) Sample imaging using microfabricated AFM cantilever with sharp tip on its end. (H) Processed image of amyloid fibrils. <a href="https://www.jove.com/files/ftp_upload/60108/60108fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60108/60108fig03.jpg" /> 
Figure 3: AFM image processing procedure42. 
The top of each panel shows a profile line of the sample surface (red line) illustrated by the corresponding AFM image, while the lower part shows a histogram of the height of all pixels in the image. (a) Raw AFM image before image flattening procedure. (b) AFM image after the processing procedure using the whole plane flattening. Fibrillar structures (pink color) were masked from the flattening procedure. (c) Image processed using line-by-line flattening procedure. (d) Final image after the image processing procedure. This figure has been adapted from Ruggeri et al.42.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60108/60108fig04.jpg" /> 
Figure 4: Single aggregate statistical analysis of AFM images. 
(a) Example of the tracing of the heights and lengths of fibrillar structures, indicated with 1 and 2. (b) Graph with sections of the traced fibrils and their average height. (c) Example of a histogram showing the average height of fibrillar structures. (d) Graph with normalized profile of the traced fibrils 1 and 2. (e) Histogram distribution of the normalized profile height points. This figure has been adapted from Ruggeri et al.42. <a href="https://www.jove.com/files/ftp_upload/60108/60108fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/60108/60108fig05.jpg" /> 
Figure 5: Principle of function of the AFM-IR method. 
(a) Absorbed IR light causes the thermal expansion of the sample, exciting the mechanical resonances of the AFM cantilever in contact with the sample. The amplitude of the cantilever oscillations is proportional to the IR absorption. (b) IR absorption maps are obtained scanning the cantilever on the sample while fixing the laser wavelength. (c) Tip and sample in contact behave as a system of coupled springs whose resonant frequency increases monotonically with the intrinsic stiffness of the sample50. (d) Localized spectra are obtained by sweeping the laser wavelength while fixing the position of the AFM cantilever. (e) IR spectrum of protein. (f) In summary, AFM-IR enables the simultaneous study of morphological, mechanical and chemical properties at the nanoscale26. <a href="https://www.jove.com/files/ftp_upload/60108/60108fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/60108/60108fig06.jpg" /> 
Figure 6: Infrared nanospectroscopy (nano-IR) of a single amyloid aggregate. 
(a) AFM morphology map. (b) IR absorption map at the laser resonance peak at 1658 cm-1. (c) Cross sectional dimensions of the fibril height. (d) IR spectra on different positions of the fibrillar structure (marked in panel b with blue circles) and the substrate (marked in panel b with green circles). The average net signal deriving from the aggregate structure (solid black line) was obtained by subtracting the averaged background signal (solid green line) from the averaged fibril signal (solid blue line). This figure has been adapted from Ruggeri et al.42.

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Characterizing Individual Protein Aggregates by Infrared Nanospectroscopy and Atomic Force Microscopy

We describe the application of infrared nanospectroscopy and high-resolution atomic force microscopy to visualize the process of protein self-assembly into oligomeric aggregates and amyloid fibrils, which is closely associated with the onset and development of a wide range of human neurodegenerative disorders.

The phenomenon of protein misfolding and aggregation results in the formation of highly heterogeneous protein aggregates, which are associated with ...

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Biochemistry

9.7K Views.  University of Cambridge. We describe the application of infrared nanospectroscopy and high-resolution atomic force microscopy to visualize the process of protein self-assembly into oligomeric aggregates and amyloid fibrils, which is closely associated with the onset and development of a wide range of human neurodegenerative disorders.

Video: Characterizing Individual Protein Aggregates by Infrared Nanospectroscopy and Atomic Force Microscopy

Enrichment of Detergent-insoluble Protein Aggregates from Human Postmortem Brain

In this study, we describe an abbreviated single-step fractionation protocol for the enrichment of detergent-insoluble protein aggregates from human postmortem brain. The ionic detergent N-lauryl-sarcosine (sarkosyl) effectively solubilizes natively folded proteins in brain tissue allowing the enrichment of detergent-insoluble protein aggregates from a wide range of neurodegenerative proteinopathies, such as Alzheimer&#39;s disease (AD), Parkinson&#39;s disease and amyotrophic lateral sclerosis, and prion diseases. Human control and AD postmortem brain tissues were homogenized and sedimented by ultracentrifugation in the presence of sarkosyl to enrich detergent-insoluble protein aggregates including pathologic phosphorylated tau, the core component of neurofibrillary tangles in AD. Western blotting demonstrated the differential solubility of aggregated phosphorylated-tau and the detergent-soluble protein, Early Endosome Antigen 1 (EEA1) in control and AD brain. Proteomic analysis also revealed enrichment of &#946;-amyloid (A&#946;), tau, snRNP70 (U1-70K), and apolipoprotein E (APOE) in the sarkosyl-insoluble fractions of AD brain compared to those of control, consistent with previous tissue fractionation strategies. Thus, this simple enrichment protocol is ideal for a wide range of experimental applications ranging from Western blotting and functional protein co-aggregation assays to mass spectrometry-based proteomics.

An abbreviated fractionation protocol for the enrichment of detergent-insoluble protein aggregates from human postmortem brain is described.

In this study, we describe an abbreviated single-step fractionation protocol for the enrichment of detergent-insoluble protein aggregates from human ...

Protein Film Infrared Electrochemistry Demonstrated for Study of H2 Oxidation by a [NiFe] Hydrogenase

Understanding the chemistry of redox proteins demands methods that provide precise control over redox centers within the protein. The technique of protein film electrochemistry, in which a protein is immobilized on an electrode surface such that the electrode replaces physiological electron donors or acceptors, has provided functional insight into the redox reactions of a range of different proteins. Full chemical understanding requires electrochemical control to be combined with other techniques that can add additional structural and mechanistic insight. Here we demonstrate a technique, protein film infrared electrochemistry, which combines protein film electrochemistry with infrared spectroscopic sampling of redox proteins. The technique uses a multiple-reflection attenuated total reflectance geometry to probe a redox protein immobilized on a high surface area carbon black electrode. Incorporation of this electrode into a flow cell allows solution pH or solute concentrations to be changed during measurements. This is particularly powerful in addressing redox enzymes, where rapid catalytic turnover can be sustained and controlled at the electrode allowing spectroscopic observation of long-lived intermediate species in the catalytic mechanism. We demonstrate the technique with experiments on E. coli hydrogenase 1 under turnover (H2 oxidation) and non-turnover conditions.

Protein Film Infrared Electrochemistry Demonstrated for Study of H2 Oxidation by a [NiFe] Hydrogenase

Here, we describe a technique, protein film infrared electrochemistry, which allows immobilized redox proteins to be studied spectroscopically under direct electrochemical control at a carbon electrode. Infrared spectra of a single protein sample can be recorded at a range of applied potentials and under a variety of solution conditions.

Understanding the chemistry of redox proteins demands methods that provide precise control over redox centers within the protein. The technique of protein ...

Force Spectroscopy of Single Protein Molecules Using an Atomic Force Microscope

The determination of the folding process of proteins from their amino acid sequence to their native 3D structure is an important problem in biology. Atomic force microscopy (AFM) can address this problem by enabling stretching and relaxation of single protein molecules, which gives direct evidence of specific unfolding and refolding characteristics. AFM-based single-molecule force-spectroscopy (AFM-SMFS) provides a means to consistently measure high-energy conformations in proteins that are not possible in traditional bulk (biochemical) measurements. Although numerous papers were published to show principles of AFM-SMFS, it is not easy to conduct SMFS experiments due to a lack of an exhaustively complete protocol. In this study, we briefly illustrate the principles of AFM and extensively detail the protocols, procedures, and data analysis as a guideline to achieve good results from SMFS experiments. We demonstrate representative SMFS results of single protein mechanical unfolding measurements and we provide troubleshooting strategies for some commonly encountered problems.

We describe the detailed procedures and strategies to measure the mechanical properties and mechanical unfolding pathways of single protein molecules using an atomic force microscope. We also show representative results as a reference for selection and justification of good single protein molecule recordings.

The determination of the folding process of proteins from their amino acid sequence to their native 3D structure is an important problem in biology. ...

Covalent Immobilization of Proteins for the Single Molecule Force Spectroscopy

In recent years, atomic force microscopy (AFM) based single molecule force spectroscopy (SMFS) extended our understanding of molecular properties and functions. It gave us the opportunity to explore a multiplicity of biophysical mechanisms, e.g., how bacterial adhesins bind to host surface receptors in more detail. Among other factors, the success of SMFS experiments depends on the functional and native immobilization of the biomolecules of interest on solid surfaces and AFM tips. Here, we describe a straightforward protocol for the covalent coupling of proteins to silicon surfaces using silane-PEG-carboxyls and the well-established N-hydroxysuccinimid/1-ethyl-3-(3-dimethyl-aminopropyl)carbodiimid (EDC/NHS) chemistry in order to explore the interaction of pilus-1 adhesin RrgA from the Gram-positive bacterium Streptococcus pneumoniae (S. pneumoniae) with the extracellular matrix protein fibronectin (Fn). Our results show that the surface functionalization leads to a homogenous distribution of Fn on the glass surface and to an appropriate concentration of RrgA on the AFM cantilever tip, apparent by the target value of up to 20% of interaction events during SMFS measurements and revealed that RrgA binds to Fn with a mean force of 52 pN. The protocol can be adjusted to couple via site specific free thiol groups. This results in a predefined protein or molecule orientation and is suitable for other biophysical applications besides the SMFS.

This protocol describes the covalent immobilization of proteins with a heterobifunctional silane coupling agent to silicon-oxide surfaces designed for the atomic force microscopy based single molecule force spectroscopy which is exemplified by the interaction of RrgA (pilus-1 tip adhesin of S. pneumoniae) with fibronectin.

In recent years, atomic force microscopy (AFM) based single molecule force spectroscopy (SMFS) extended our understanding of molecular properties and ...

Analysis of &#946;-Amyloid-induced Abnormalities on Fibrin Clot Structure by Spectroscopy and Scanning Electron Microscopy

This article presents methods for generating in vitro fibrin clots and analyzing the effect of beta-amyloid (A&#946;) protein on clot formation and structure by spectrometry and scanning electron microscopy (SEM). A&#946;, which forms neurotoxic amyloid aggregates in Alzheimer&#39;s disease (AD), has been shown to interact with fibrinogen. This A&#946;-fibrinogen interaction makes the fibrin clot structurally abnormal and resistant to fibrinolysis. A&#946;-induced abnormalities in fibrin clotting may also contribute to cerebrovascular aspects of the AD pathology such as microinfarcts, inflammation, as well as, cerebral amyloid angiopathy (CAA). Given the potentially critical role of neurovascular deficits in AD pathology, developing compounds which can inhibit or lessen the A&#946;-fibrinogen interaction has promising therapeutic value. In vitro methods by which fibrin clot formation can be easily and systematically assessed are potentially useful tools for developing therapeutic compounds. Presented here is an optimized protocol for in vitro generation of the fibrin clot, as well as analysis of the effect of A&#946; and A&#946;-fibrinogen interaction inhibitors. The clot turbidity assay is rapid, highly reproducible and can be used to test multiple conditions simultaneously, allowing for the screening of large numbers of A&#946;-fibrinogen inhibitors. Hit compounds from this screening can be further evaluated for their ability to ameliorate A&#946;-induced structural abnormalities of the fibrin clot architecture using SEM. The effectiveness of these optimized protocols is demonstrated here using TDI-2760, a recently identified A&#946;-fibrinogen interaction inhibitor.

Presented here are two methods that can be used individually or in combination to analyze the effect of beta-amyloid on fibrin clot structure. Included is a protocol for creating an in vitro fibrin clot, followed by clot turbidity and scanning electron microscopy methods.

This article presents methods for generating in vitro fibrin clots and analyzing the effect of beta-amyloid (A&#946;) protein on clot formation and ...

High Sensitivity Measurement of Transcription Factor-DNA Binding Affinities by Competitive Titration Using Fluorescence Microscopy

Accurate quantification of transcription factor (TF)-DNA interactions is essential for understanding the regulation of gene expression. Since existing approaches suffer from significant limitations, we have developed a new method for determining TF-DNA binding affinities with high sensitivity on a large scale. The assay relies on the established fluorescence anisotropy (FA) principle but introduces important technical improvements. First, we measure a full FA competitive titration curve in a single well by incorporating TF and a fluorescently labeled reference DNA in a porous agarose gel matrix. Unlabeled DNA oligomer is loaded on the top as a competitor and, through diffusion, forms a spatio-temporal gradient. The resulting FA gradient is then read out using a customized epifluorescence microscope setup. This improved setup greatly increases the sensitivity of FA signal detection, allowing both weak and strong binding to be reliably quantified, even for molecules of similar molecular weights. In this fashion, we can measure one titration curve per well of a multi-well plate, and through a fitting procedure, we can extract both the absolute dissociation constant (KD) and active protein concentration. By testing all single-point mutation variants of a given consensus binding sequence, we can survey the entire binding specificity landscape of a TF, typically on a single plate. The resulting position weight matrices (PWMs) outperform those derived from other methods in predicting in vivo TF occupancy. Here, we present a detailed guide for implementing HiP-FA on a conventional automated fluorescent microscope and the data analysis pipeline.

Here we present a novel method for determining binding affinities at equilibrium and in solution with high sensitivity on a large scale. This improves the quantitative analysis of transcription factor-DNA binding. The method is based on automated fluorescence anisotropy measurements in a controlled delivery system.

Accurate quantification of transcription factor (TF)-DNA interactions is essential for understanding the regulation of gene expression. Since existing ...

Probing The Structure And Dynamics Of Nucleosomes Using Atomic Force Microscopy Imaging

Chromatin, which is a long chain of nucleosome subunits, is a dynamic system that allows for such critical processes as DNA replication and transcription to take place in eukaryotic cells. The dynamics of nucleosomes provides access to the DNA by replication and transcription machineries, and critically contributes to the molecular mechanisms underlying chromatin functions. Single-molecule studies such as atomic force microscopy (AFM) imaging have contributed significantly to our current understanding of the role of nucleosome structure and dynamics. The current protocol describes the steps enabling high-resolution AFM imaging techniques to study the structural and dynamic properties of nucleosomes. The protocol is illustrated by AFM data obtained for the centromere nucleosomes in which H3 histone is replaced with its counterpart centromere protein A (CENP-A). The protocol starts with the assembly of mono-nucleosomes using a continuous dilution method. The preparation of the mica substrate functionalized with aminopropyl silatrane (APS-mica) that is used for the nucleosome imaging is critical for the AFM visualization of nucleosomes described and the procedure to prepare the substrate is provided. Nucleosomes deposited on the APS-mica surface are first imaged using static AFM, which captures a snapshot of the nucleosome population. From analyses of these images, such parameters as the size of DNA wrapped around the nucleosomes can be measured and this process is also detailed. The time-lapse AFM imaging procedure in the liquid is described for the high-speed time-lapse AFM that can capture several frames of nucleosome dynamics per second. Finally, the analysis of nucleosome dynamics enabling the quantitative characterization of the dynamic processes is described and illustrated.

Here, we present a protocol to characterize nucleosome particles at the single-molecule level using static and time-lapse atomic force microscopy (AFM) imaging techniques. The surface functionalization method described allows for the capture of the structure and dynamics of nucleosomes in high-resolution at the nanoscale.

Chromatin, which is a long chain of nucleosome subunits, is a dynamic system that allows for such critical processes as DNA replication and transcription ...

Analysis of Protein Folding, Transport, and Degradation in Living Cells by Radioactive Pulse Chase

Radioactive pulse-chase labeling is a powerful tool for studying the conformational maturation, the transport to their functional cellular location, and the degradation of target proteins in live cells. By using short (pulse) radiolabeling times (&lt;30 min) and tightly controlled chase times, it is possible to label only a small fraction of the total protein pool and follow its folding. When combined with nonreducing/reducing SDS-polyacrylamide gel electrophoresis (SDS-PAGE) and immunoprecipitation with (conformation-specific) antibodies, folding processes can be examined in great detail. This system has been used to analyze the folding of proteins with a huge variation in properties such as soluble proteins, single and multi-pass transmembrane proteins, heavily N- and O-glycosylated proteins, and proteins with and without extensive disulfide bonding. Pulse-chase methods are the basis of kinetic studies into a range of additional features, including co- and posttranslational modifications, oligomerization, and polymerization, essentially allowing the analysis of a protein from birth to death. Pulse-chase studies on protein folding are complementary with other biochemical and biophysical methods for studying proteins in vitro by providing increased temporal resolution and physiological information. The methods as described within this paper are adapted easily to study the folding of almost any protein that can be expressed in mammalian or insect-cell systems.

Here we describe a protocol for a general pulse-chase method that allows the kinetic analysis of folding, transport, and degradation of proteins to be followed in live cells.

Radioactive pulse-chase labeling is a powerful tool for studying the conformational maturation, the transport to their functional cellular location, and ...

Imaging of Extracellular Vesicles by Atomic Force Microscopy

Exosomes and other extracellular vesicles (EVs) are molecular complexes consisting of a lipid membrane vesicle, its surface decoration by membrane proteins and other molecules, and diverse luminal content inherited from a parent cell, which includes RNAs, proteins, and DNAs. The characterization of the hydrodynamic sizes of EVs, which depends on the size of the vesicle and its coronal layer formed by surface decorations, has become routine. For exosomes, the smallest of EVs, the relative difference between the hydrodynamic and vesicles sizes is significant. The characterization of vesicles sizes by the cryogenic transmission electron microscopy (cryo-TEM) imaging, a gold standard technique, remains a challenge due to the cost of the instrument, the expertise required to perform the sample preparation, imaging and data analysis, and a small number of particles often observed in images. A widely available and accessible alternative is the atomic force microscopy (AFM), which can produce versatile data on three-dimensional geometry, size, and other biophysical properties of extracellular vesicles. The developed protocol guides the users in utilizing this analytical tool and outlines the workflow for the analysis of EVs by the AFM, which includes the sample preparation for imaging EVs in hydrated or desiccated form, the electrostatic immobilization of vesicles on a substrate, data acquisition, its analysis, and interpretation. The representative results demonstrate that the fixation of EVs on the modified mica surface is predictable, customizable, and allows the user to obtain sizing results for a large number of vesicles. The vesicle sizing based on the AFM data was found to be consistent with the cryo-TEM imaging.&#160;

A step-by-step procedure is described for label-free immobilization of exosomes and extracellular vesicles from liquid samples and their imaging by atomic force microscopy (AFM). The AFM images are used to estimate the size of the vesicles in the solution and characterize other biophysical properties.&#160;

Exosomes and other extracellular vesicles (EVs) are molecular complexes consisting of a lipid membrane vesicle, its surface decoration by membrane ...

Fully Processed Recombinant KRAS4b: Isolating and Characterizing the Farnesylated and Methylated Protein

Protein prenylation is a key modification that is responsible for targeting proteins to intracellular membranes. KRAS4b, which is mutated in 22% of human cancers, is processed by farnesylation and carboxymethylation due to the presence of a &#39;CAAX&#39; box motif at the C-terminus. An engineered baculovirus system was used to express farnesylated and carboxymethylated KRAS4b in insect cells and has been described previously. Here, we describe the detailed, practical purification and biochemical characterization of the protein. Specifically, affinity and ion exchange chromatography were used to purify the protein to homogeneity. Intact and native mass spectrometry was used to validate the correct modification of KRAS4b and to verify nucleotide binding. Finally, membrane association of farnesylated and carboxymethylated KRAS4b to liposomes was measured using surface plasmon resonance spectroscopy.

Prenylation is an important modification on peripheral membrane binding proteins. Insect cells can be manipulated to produce farnesylated and carboxymethylated KRAS4b in quantities that enable biophysical measurements of protein-protein and protein-lipid interactions

Protein prenylation is a key modification that is responsible for targeting proteins to intracellular membranes. KRAS4b, which is mutated in 22% of human ...