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 ...

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 optimize...

Watch this Scientific Journal Video about Characterizing Individual Protein Aggregates by Infrared Nanospectroscopy and Atomic Force Microscopy at JoVE.com

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 ...

characterizing-individual-protein-aggregates-infrared

Universidad San Sebastian

Research

JoVE Journal

Biochemistry

9.7K Views.  University of Cambridge. Methoden op basis van atomaire kracht microscopie maken het mogelijk om biomoleculaire processen op nanoschaal te karakteriseren met morfologische chemische of structurele resolutie, eindelijk het openen van een nieuwe voorste venster van observatie op de biologie. Methoden van één molecuul maken indringende zeer heterogene systemen mogelijk, een kenmerk dat bijzonder relevant is voor het begrijpen van eiwitaggregatie. Door het combineren van enkele moleculen met rugbenaderingen, kunnen we ...

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

Visualization of Recombinant DNA and Protein Complexes Using Atomic Force Microscopy

Atomic force microscopy (AFM) allows for the visualizing of individual proteins, DNA molecules, protein-protein complexes, and DNA-protein complexes. On the end of the microscope's cantilever is a nano-scale probe, which traverses image areas ranging from nanometers to micrometers, measuring the elevation of macromolecules resting on the substrate surface at any given point. Electrostatic forces cause proteins, lipids, and nucleic acids to loosely attach to the substrate in random orientations and permit imaging. The generated data resemble a topographical map, where the macromolecules resolve as three-dimensional particles of discrete sizes (Figure 1) 1,2. Tapping mode AFM involves the repeated oscillation of the cantilever, which permits imaging of relatively soft biomaterials such as DNA and proteins. One of the notable benefits of AFM over other nanoscale microscopy techniques is its relative adaptability to visualize individual proteins and macromolecular complexes in aqueous buffers, including near-physiologic buffered conditions, in real-time, and without staining or coating the sample to be imaged. 
The method presented here describes the imaging of DNA and an immunoadsorbed transcription factor (i.e. the glucocorticoid receptor, GR) in buffered solution (Figure 2). Immunoadsorbed proteins and protein complexes can be separated from the immunoadsorbing antibody-bead pellet by competition with the antibody epitope and then imaged (Figure 2A). This allows for biochemical manipulation of the biomolecules of interest prior to imaging. Once purified, DNA and proteins can be mixed and the resultant interacting complex can be imaged as well. Binding of DNA to mica requires a divalent cation 3,such as Ni2+ or Mg2+, which can be added to sample buffers yet maintain protein activity. Using a similar approach, AFM has been utilized to visualize individual enzymes, including RNA polymerase 4 and a repair enzyme 5, bound to individual DNA strands. These experiments provide significant insight into the protein-protein and DNA-protein biophysical interactions taking place at the molecular level. Imaging individual macromolecular particles with AFM can be useful for determining particle homogeneity and for identifying the physical arrangement of constituent components of the imaged particles. While the present method was developed for visualization of GR-chaperone protein complexes 1,2 and DNA strands to which the GR can bind, it can be applied broadly to imaging DNA and protein samples from a variety of sources.

A tapping mode atomic force microscope (AFM) method for the visualization of plasmid DNA, cytoplasmic proteins, and DNA-protein complexes is described. The method includes alternate approaches for preparing samples for AFM imaging following biochemical manipulation. DNA containing specific protein interacting regions are observed in near-physiologic buffer conditions.

Atomic force microscopy (AFM) allows for the visualizing of individual proteins, DNA molecules, protein-protein complexes, and DNA-protein complexes. On ...

Bioengineering

Atomic Force Microscopy of Red-Light Photoreceptors Using PeakForce Quantitative Nanomechanical Property Mapping

Atomic force microscopy (AFM) uses a pyramidal tip attached to a cantilever to probe the force response of a surface. The deflections of the tip can be measured to ~10 pN by a laser and sectored detector, which can be converted to image topography. Amplitude modulation or &#8220;tapping mode&#8221; AFM involves the probe making intermittent contact with the surface while oscillating at its resonant frequency to produce an image. Used in conjunction with a fluid cell, tapping-mode AFM enables the imaging of biological macromolecules such as proteins in physiologically relevant conditions. Tapping-mode AFM requires manual tuning of the probe and frequent adjustments of a multitude of scanning parameters which can be challenging for inexperienced users. To obtain high-quality images, these adjustments are the most time consuming.
PeakForce Quantitative Nanomechanical Property Mapping (PF-QNM) produces an image by measuring a force response curve for every point of contact with the sample. With ScanAsyst software, PF-QNM can be automated. This software adjusts the set-point, drive frequency, scan rate, gains, and other important scanning parameters automatically for a given sample. Not only does this process protect both fragile probes and samples, it significantly reduces the time required to obtain high resolution images. PF-QNM is compatible for AFM imaging in fluid; therefore, it has extensive application for imaging biologically relevant materials.
The method presented in this paper describes the application of PF-QNM to obtain images of a bacterial red-light photoreceptor, RpBphP3 (P3), from photosynthetic R. palustris in its light-adapted state. Using this method, individual protein dimers of P3 and aggregates of dimers have been observed on a mica surface in the presence of an imaging buffer. With appropriate adjustments to surface and/or solution concentration, this method may be generally applied to other biologically relevant macromolecules and soft materials.

A method for investigating the structure of a protein photoreceptor using atomic force microscopy (AFM) is described in this paper. PeakForce Quantitative Nanomechanical Property Mapping (PF-QNM) reveals intact protein dimers on a mica surface.

Atomic force microscopy (AFM) uses a pyramidal tip attached to a cantilever to probe the force response of a surface. The deflections of the tip can be ...

Engineering

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 ...

OaAEP1-Mediated Enzymatic Synthesis and Immobilization of Polymerized Protein for Single-Molecule Force Spectroscopy

Chemical and bio-conjugation techniques have been developed rapidly in recent years and allow the building of protein polymers. However, a controlled protein polymerization process is always a challenge. Here, we have developed an enzymatic methodology for constructing polymerized protein step by step in a rationally-controlled sequence. In this method, the C-terminus of a protein monomer is NGL for protein conjugation using OaAEP1 (Oldenlandia affinis asparaginyl endopeptidases) 1) while the N-terminus was a cleavable TEV (tobacco etch virus) cleavage site plus an L (ENLYFQ/GL) for temporary N-terminal protecting. Consequently, OaAEP1 was able to add only one protein monomer at a time, and then the TEV protease cleaved the N-terminus between Q and G to expose the NH2-Gly-Leu. Then the unit is ready for next OaAEP1 ligation. The engineered polyprotein is examined by unfolding individual protein domain using atomic force microscopy-based single-molecule force spectroscopy (AFM-SMFS). Therefore, this study provides a useful strategy for polyprotein engineering and immobilization.

Here, we present a protocol to conjugate protein monomer by enzymes forming protein polymer with a controlled sequence and immobilize it on the surface for single-molecule force spectroscopy studies.

Chemical and bio-conjugation techniques have been developed rapidly in recent years and allow the building of protein polymers. However, a controlled ...

Assembly, Tuning and Use of an Apertureless Near Field Infrared Microscope for Protein Imaging

This paper aims to instruct the reader in the assembly and operation of an infrared near-field microscope for imaging beyond the diffraction limit.  The apertureless near-field microscope is a light scattering-type instrument that provides infrared spectra at circa 20 nm resolution. A complete list of components and a step-by-step protocol for use is provided.  Common errors in assembly and instrument tuning are discussed.  A representative data set that shows the secondary structure of an amyloid fibril is presented.  

The assembly of a nearfield infrared microscope for imaging protein aggregates is described.

This paper aims to instruct the reader in the assembly and operation of an infrared near-field microscope for imaging beyond the diffraction limit.  The ...

Biology

Atomic Force Microscopy Combined with Infrared Spectroscopy as a Tool to Probe Single Bacterium Chemistry

Atomic Force Microscopy-Infrared Spectroscopy (AFM-IR) is a novel combinatory technique, enabling simultaneous characterization of physical properties and chemical composition of sample with nanoscale resolution. By combining AFM with IR, the spatial resolution limitation of conventional IR is overcome, enabling a resolution of 20&#8211;100 nm to be achieved. This opens the door for a broad array of new applications of IR toward probing samples smaller than several micrometers, previously unachievable by means of conventional IR microscopy. AFM-IR is eminently suited for bacterial research, providing both spectral and spatial information at the single cell and intracellular level. The increasing global health concerns and unfavorable future prediction regarding bacterial infections, and especially, rapid development of antimicrobial resistance, has created an urgent need for a research tool capable of phenotypic probing at the single cell and subcellular level. AFM-IR offers the potential to address this need, by enabling detail characterization of chemical composition of a single bacterium. Here, we provide a complete protocol for sample preparation and data acquisition of single spectra and mapping modality, for the application of AFM-IR toward bacterial studies.

Atomic Force Microscopy-Infrared Spectroscopy (AFM-IR) provides a powerful platform for bacterial studies, enabling to achieve nanoscale resolution. Both, mapping of subcellular changes (e.g., upon cell division) as well as comparative studies of chemical composition (e.g., arising from drug resistance) can be conducted at a single cell level in bacteria.

Atomic Force Microscopy-Infrared Spectroscopy (AFM-IR) is a novel combinatory technique, enabling simultaneous characterization of physical properties and ...

Chemistry

High-Resolution Neutron Spectroscopy to Study Picosecond-Nanosecond Dynamics of Proteins and Hydration Water

Neutron scattering offers the possibility to probe the dynamics within samples for a wide range of energies in a nondestructive manner and without labeling other than deuterium. In particular, neutron backscattering spectroscopy records the scattering signals at multiple scattering angles simultaneously and is well suited to study the dynamics of biological systems on the ps-ns timescale. By employing D2O-and possibly deuterated buffer components-the method allows monitoring of both center-of-mass diffusion and backbone and side-chain motions (internal dynamics) of proteins in liquid state.
Additionally, hydration water dynamics can be studied by employing powders of perdeuterated proteins hydrated with H2O. This paper presents the workflow employed on the instrument IN16B at the Institut Laue-Langevin (ILL) to investigate protein and hydration water dynamics. The preparation of solution samples and hydrated protein powder samples using vapor exchange is explained. The data analysis procedure for both protein and hydration water dynamics is described for different types of datasets (quasielastic spectra or fixed-window scans) that can be obtained on a neutron backscattering spectrometer.
The method is illustrated with two studies involving amyloid proteins. The aggregation of lysozyme into &#181;m sized spherical aggregates-denoted particulates-is shown to occur in a one-step process on the space and time range probed on IN16B, while the internal dynamics remains unchanged. Further, the dynamics of hydration water of tau was studied on hydrated powders of perdeuterated protein. It is shown that translational motions of water are activated upon the formation of amyloid fibers. Finally, critical steps in the protocol are discussed as to how neutron scattering is positioned regarding the study of dynamics with respect to other experimental biophysical methods.

Neutron backscattering spectroscopy offers a nondestructive and label-free access to the ps-ns dynamics of proteins and their hydration water. The workflow is presented with two studies on amyloid proteins: on the time-resolved dynamics of lysozyme during aggregation and on the hydration water dynamics of tau upon fiber formation.

Neutron scattering offers the possibility to probe the dynamics within samples for a wide range of energies in a nondestructive manner and without ...

Exploring Multiphase Polymeric Systems by Nanoscale Infrared Spectroscopy

Advances in Nanoscale Infrared Spectroscopy to Explore Multiphase Polymeric Systems

Multiphase polymeric systems encompass local domains with dimensions that can vary from a few tens of nanometers to several micrometers. Their composition is commonly assessed using infrared spectroscopy, which provides an average fingerprint of the various materials contained in the volume probed. However, this approach does not offer any details on the arrangement of the phases in the material. Interfacial regions between two polymeric phases, often in the nanoscale range, are also challenging to access. Photothermal nanoscale infrared spectroscopy monitors the local response of materials excited by infrared light with the sensitive probe of an atomic force microscope (AFM). While the technique is suitable for interrogating small features, such as individual proteins on pristine gold surfaces, the characterization of three-dimensional multicomponent materials is more elusive. This is due to a relatively large volume of material undergoing photothermal expansion, defined by the laser focalization onto the sample and by the thermal properties of the polymeric constituents, compared to the nanoscale region probed by the AFM tip. Using a polystyrene (PS) bead and a polyvinyl alcohol (PVA) film, we evaluate the spatial footprint of photothermal nanoscale infrared spectroscopy for surface analysis as a function of the position of PS in the PVA film. The effect of the feature position on the nanoscale infrared images is investigated, and spectra are acquired. Some perspectives on the future advances in the field of photothermal nanoscale infrared spectroscopy are provided, considering the characterization of complex systems with embedded polymeric structures.

Author Spotlight: Advances in Nanoscale Infrared Spectroscopy to Explore Multiphase Polymeric Systems  

This protocol describes the application of atomic force microscopy and nanoscale infrared spectroscopy to evaluate the performance of photothermal nanoscale infrared spectroscopy in the characterization of three-dimensional multi-polymeric samples.

Multiphase polymeric systems encompass local domains with dimensions that can vary from a few tens of nanometers to several micrometers. Their composition ...

Detection of Protein Aggregation using Fluorescence Correlation Spectroscopy

Protein aggregation is a hallmark of neurodegenerative disorders such as amyotrophic lateral sclerosis (ALS), Alzheimer's disease (AD), Parkinson's disease (PD), Huntington's disease (HD), and so on. To detect and analyze soluble or diffuse protein oligomers or aggregates, fluorescence correlation spectroscopy (FCS), which can detect the diffusion speed and brightness of a single particle with a single molecule sensitivity, has been used. However, the proper procedure and know-how for protein aggregation detection have not been widely shared. Here, we show a standard procedure of FCS measurement for diffusion properties of aggregation-prone proteins in cell lysate and live cells: ALS-associated 25 kDa carboxyl-terminal fragment of TAR DNA/RNA-binding protein 43 kDa (TDP25) and superoxide dismutase 1 (SOD1). The representative results show that a part of aggregates of green fluorescent protein (GFP)-tagged TDP25 was slightly included in the soluble fraction of murine neuroblastoma Neuro2a cell lysate. Moreover, GFP-tagged SOD1 carrying ALS-associated mutation shows a slower diffusion in live cells. Accordingly, we here introduce the procedure to detect the protein aggregation via its diffusion property using FCS.

We here introduce a procedure to measure protein oligomers and aggregation in cell lysate and live cells using fluorescence correlation spectroscopy.

Protein aggregation is a hallmark of neurodegenerative disorders such as amyotrophic lateral sclerosis (ALS), Alzheimer's disease (AD), Parkinson's ...

Characterizing Individual Protein Aggregates by Infrared Nanospectroscopy and Atomic Force Microscopy

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