Name: characterizing individual protein aggregates by infrared nanospectroscopy and atomic force microscopy
Uploaded: 2019-09-12T15:00:00
Description: Watch this Scientific Journal Video about Characterizing Individual Protein Aggregates by Infrared Nanospectroscopy and Atomic Force Microscopy at JoVE.com

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 aggregatio...

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 ...

Methods based on atomic force microscopy make it possible to characterize biomolecular processes at the nano scale with morphological chemical or structural resolution, finally opening a new front window of observation on biology. Single molecule methods enable probing highly heterogeneous systems, a feature which is particularly relevant for understanding protein aggregation. By combining single molecules with back approaches, we can obtain fundamental information about the behavior of proteins, their links with human disease and design pharmacological interventions which can be successful.Furthermore, this method can be successfully applied to study the chemical and structural properties of biomaterials for drug delivery, applications and other biotechnological purposes. It may be challenging initially to set the correct AFM parameters for high resolution and it's easy to set up your AFM while measuring your fibrillar aggregates. 30 minutes before the AFM imaging, turn on the AFM system and click set up and probe.In the probe change window, click next to prepare Z focus stage. Turn off the AFM beam switch if it is on and unlock the dovetail locks. Disconnect the head from the system and in the probe change window, click next.Mount the AFM cantilever on the probe holder. Connect the head to the system and lock the dovetail locks. Turn on the AFM laser beam switch and in the probe change window, click next.In the pop up window, click no if the cantilever type did not change and adjust the optical alignment knobs to find the cantilever. Adjust the focus onto the cantilever and click next. In the pop up window, click yes if the focus is on the cantilever.Use the detection laser beam knobs to position the laser beam at the end of the cantilever. Use the deflected laser beam knobs to maximize the total signal measured by the four quadrant photo diode to at least greater than one volt. In the probe change window, click next and close.After waiting 15 minutes for the cantilever to reach thermal stability, readjust the position of the deflected laser beam on the position sensitive photo diode if necessary. Click NCM sweep, select the desired amplitude of oscillation, click use phase and click auto. Tune the cantilever close to the maximum of its first free resonance of oscillation to approximately 300 kilohertz for a cantilever with a spring constant of 40 newtons per meter.Place the sample on the sample holder. Click scan area and set the resolution to between 256 by 256 and 1024 by 1024 pixels and then set the scan size and the pixel number. Set the scanning rate to 0.3 to one hertz for a scan area of one by one to five by five micrometers squared.Focus the optical view on the sample and click approach to approach the sample surface. Next, click lift 100 micrometers to raise the AFM tip 100 micrometers above the surface of the sample and click expand on the optical image of the sample. Click the focus stage bar to focus the view on the surface of the sample and use the arrows to move in the region of the sample of interest.Click approach to engage the surface and click line scan to check if the tip is following the surface well. Adjust the set point as necessary. Then click scan to begin imaging the sample surface, maintaining a constant regime of phase change not exceeding delta 20 during the imaging to avoid a large imaging force and to maintain consistency between independent samples.For IR nanospectroscopy imaging, 30 to 60 minutes before the analysis, turn on the AFM IR system and the IR laser. Open the built in software to control the instrument and click file and new to open a new nano IR file. Click initialize to start the AFM IR system and open the instrument cover.Mount a silicone gold coated probe with a nominal radius of 30 nanometers and a spring constant of 0.2 newtons per meter onto the AFM IR system to measure the sample in contact mode. Click load in the AFM probe section and next. Use the focus on probe arrows to focus the camera on the cantilever and use the tip to crosshairs to place the crosshair at the end of the cantilever.Rotate the knobs controlling the position of the detection laser to position the laser at the end of the cantilever. Rotate the laser knobs to detect and maximize the sum measured by the four quadrant photo diode to a value greater than three volts. Rotate the deflection knob to adjust the cantilever deflection to minus one volt and click next.Then close the cover of the instrument. Use the focus on probe arrows to focus the camera on the sample and tip to crosshairs arrows to move in the region of interest of the sample. Click next and engage.In the microscope window, select height for morphology, amplitude two for the IR absorption and PLL frequency to map the tip to sample contact resonance. In the AFM scan section, set the parameters as demonstrated for the AFM imaging and click scan to acquire a morphology map. When the morphology mapping is finished, in the microscope window, click the height map to position the probe on the top of one aggregate.In the nano IR section, click start IR to illuminate the sample with the IR laser. To focus the infrared laser on the cantilever, click optimize and enter 1655 centimeters for the wavenumber. Click add and scan to determine the IR laser position and click update and okay to align its position with the cantilever.In the general section, enter a wavenumber for which a high absorbance in the relative field is expected and deactivate the band pass filter option. Click start IR then check the meter reading and the 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 200 kilohertz. Enter the generated resonance frequency value in the general section in the frequency center field and use a frequency window of 50 kilohertz. Click laser pulse tune window to select the resonance enhanced mode and set a pulse rate of 222 kilohertz, a tune range of 50 kilohertz and a laser duty cycle of 5%Click acquire to sweep the pulse rate of the laser.Use the cursor to tune the laser pulse to the frequency of the mechanical response of the thermal expansion of the sample absorbing the IR light. Then select phase locked loop to monitor the contact resonance between the sample and the tip and click zero. For other chemical properties at the nano scale, track the contact resonance during the spectra acquisition to ensure that the sample spectrum is not affected by overheating and softening.Select enable to track the sample to tip contact resonance and select an integral gain of 0.5 and a proportional gain of 10 then click OK.In the optimize window, click the wavelength and scan to locate the IR laser for at least three wavenumbers corresponding to major absorbance bands of the sample and for at least one wavenumber for each chip of the laser. Click tools, IR background calibration and new, and set the wavenumbers to between 1200 and 1800 centimeters. Set the backgrounds to average to one, the duty cycle of 5%and the sweep speed to 100 centimeters squared.Click acquire to measure the IR laser background for normalization of the measured nano scale localized spectra, save the file and close the window. In the IR spectra settings, select an IR spectrum resolution between one and four centimeters and a number of co-averages of at least 64 times. Then click acquire to measure a nano scale localized IR spectrum in the protein range.To acquire a nano scale resolved chemical map, select 1655 centimeters and IR imaging and click scan in the AFM scan window. When the mapping is completed, save the measurements. Then use the built in AFM image processing software to analyze the acquired maps of morphology, contact resonance and chemistry and nano scale localized spectra.It is a critical step to obtain a highly pure monomeric solution as the presence of aggregated species may result in poor reproducibility of kinetics and introduce artifacts in your measurements. A fundamental factor for successfully studying biological samples which have high heterogeneity is the correct the position of solid substrates. Microfluidic spray deposition can be exploited instead of manual one in order to preserve the sample architecture and heterogeneity.These methods pave the way for unraveling the chemical structure and properties of individual biomolecules and their interactions in physiological and native liquid environment.

characterizing-individual-protein-aggregates-infrared

Research

JoVE Journal

Biochemistry

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 ...

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 ...

Creating Highly Specific Chemically Induced Protein Dimerization Systems by Stepwise Phage Selection of a Combinatorial Single-Domain Antibody Library

Protein dimerization events that occur only in the presence of a small-molecule ligand enable the development of small-molecule biosensors for the dissection and manipulation of biological pathways. Currently, only a limited number of chemically induced dimerization (CID) systems exist and engineering new ones with desired sensitivity and selectivity for specific small-molecule ligands remains a challenge in the field of protein engineering. We here describe a high throughput screening method, combinatorial binders-enabled selection of CID (COMBINES-CID), for the de novo engineering of CID systems applicable to a large variety of ligands. This method uses the two-step selection of a phage-displayed combinatorial nanobody library to obtain 1) &#34;anchor binders&#34; that first bind to a ligand of interest and then 2) &#34;dimerization binders&#34; that only bind to anchor binder-ligand complexes. To select anchor binders, a combinatorial library of over 109 complementarity-determining region (CDR)-randomized nanobodies is screened with a biotinylated ligand and hits are validated with the unlabeled ligand by bio-layer interferometry (BLI). To obtain dimerization binders, the nanobody library is screened with anchor binder-ligand complexes as targets for positive screening and the unbound anchor binders for negative screening. COMBINES-CID is broadly applicable to select CID binders with other immunoglobulin, non-immunoglobulin, or computationally designed scaffolds to create biosensors for in vitro and in vivo detection of drugs, metabolites, signaling molecules, etc.

Creating chemically induced protein dimerization systems with desired affinity and specificity for any given small molecule ligand would have many biological sensing and actuation applications. Here, we describe an efficient, generalizable method for de novo engineering of chemically induced dimerization systems via the stepwise selection of a phage-displayed combinatorial single-domain antibody library.