This work was supported by the National Natural Science Foundation of China (Grant No. 21771103, 21977047), Natural Science Foundation of Jiangsu Province (Grant No. BK20160639) and Shuangchuang Program of Jiangsu Province.

1. Protein production
<ol>
	<li>Gene clone
	<ol>
		<li>Purchase genes coding for the protein of interest (POI): Ubiquitin, Rubredoxin (RD)51, the cellulose-binding module (CBM), dockerin-X domain (XDoc) and cohesion from Ruminococcus flavefacience, tobacco etch virus (TEV) protease, elastin-like polypeptides (ELPs).</li>
		<li>Perform polymerase chain reaction and use three-restriction digestion enzyme system BamHI-BglII-KpnI for recombining the gene from different protein fragments.</li>
		<li>Confirm all genes by direct DNA sequencing.</li>
	</ol>
	</li>
	<li>Proteins expression and production
	<ol>
		<li>Transform E. coli BL21(DE3) with the pQE80L-POI or pET28a-POI plasmid for expression.</li>
		<li>Pick one single colony into 15 mL of LB medium with respective antibiotics (e.g., 100 &#956;g/mL ampicillin sodium salt or 50 &#956;g/mL, kanamycin). Keep shaking the cultures at 200 rpm at 37 &#176;C for 16-20 h.</li>
		<li>Dilute the overnight cultures into 800 mL of LB medium (1:50 dilution). For rubredoxin, centrifuge the culture at 1,800 x g, then resuspend in 15 mL of M9 medium (supplemented with 0.4% glucose, 0.1 mM CaCl2, 2 mM MgSO4), and then dilute it into 800 mL of M9 medium.</li>
		<li>Incubate the culture at 37 &#176;C while shaking at 200 rpm, until the culture reaches an optical density at 600 nm (OD600) of 0.6. Save 100 &#956;L sample of the culture as the pre-induction control for testing protein expression.</li>
		<li>Induce protein expression by adding IPTG to a final concentration of 1 mM and shake the culture at 37 &#176;C for 4 h at 200 rpm. Reserve a 100 &#956;L sample of the culture as the post-induction control for testing protein expression.</li>
		<li>Centrifuge the culture at 13,000 x g for 25 min at 4 &#176;C and store at -80 &#176;C before purification. 
		NOTE: The protocol can be paused here.</li>
	</ol>
	</li>
	<li>Purification of protein of interest
	<ol>
		<li>Resuspend the cells in 25 mL of lysis buffer (50 mM Tris, 150 mM NaCl, pH 7.4 containing DNase, RNase, PMSF) and use a sonicator (15% amplitude) to lyse it for 30 min on ice.</li>
		<li>Clarify the cell lysate at 19,000 x g for 40 min at 4 &#176;C.</li>
		<li>Pack 1 mL (bed volume) of Co-NTA or Ni-NTA affinity column and wash the column with 10 column volumes (CV) of ultrapure water and then 10 CVs of wash buffer (50 mM Tris, 150 mM NaCl, 2 mM imidazole, pH 7.4) by gravity flow.</li>
		<li>Pass the protein supernatant through the column by gravity flow for three times.</li>
		<li>Pour wash buffer on the column with 50 CVs to move away contaminant proteins.</li>
		<li>Elute the bound protein with 3 CVs of ice-cold elution buffer (20 mM Tris, 400 mM NaCl, 250 mM imidazole, pH 7.4). When it comes to rubredoxin proteins, further anion exchange purification using anion exchange column at pH 8.5 at 4 &#176;C is necessary.</li>
		<li>Analyze the sample by SDS-PAGE.</li>
	</ol>
	</li>
</ol>
2. Functionalization of coverslip and cantilever surface
<ol>
	<li>Functionalized coverslip surface preparation
	<ol>
		<li>Dissolve 20 g of potassium chromate in 40 mL of ultrapure water. Slowly add 360 mL of concentrated sulfuric acid to the potassium chromate solution with glass rod stir gently and to prepare the chromic acid. 
		CAUTION: The chemical used here and the final chromic acid is strongly corrosive and acidic. Work with proper protective equipment. The solution releases heat when add concentrated sulfuric acid, which means slow adding and proper pause for cooling down.</li>
		<li>Clean and activate a glass coverslip at 80 &#176;C for 30 min by chromic acid treatment. Completely immerse the coverslips in 1% (v/v) APTES toluene solution for 1 h at room temperature while protecting them from light.</li>
		<li>Wash the coverslip with toluene and absolute ethyl alcohol and dry the coverslip with a stream of nitrogen.</li>
		<li>Incubate the coverslip at 80 &#176;C for 15 min and then cool down to room temperature.</li>
		<li>Add 200 &#956;L of sulfo-SMCC (1 mg/mL) in dimethyl sulfoxide (DMSO) solution between two immobilized coverslips and incubate for 1 h protected from light.</li>
		<li>Wash the coverslip with DMSO first and then with absolute ethyl alcohol to remove residual sulfo-SMCC.</li>
		<li>Dry the coverslip under a stream of nitrogen.</li>
		<li>Pipet 60 &#956;L of 200 &#956;M GL-ELP50nm-C protein solution onto a functionalized coverslip and incubate for about 3 h.</li>
		<li>Wash the coverslip with ultrapure water to remove the unreacted GL-ELP50nm-C. 
		NOTE: Functionalized coverslips are capable for about two weeks under storage at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Functionalized cantilever surface preparation
	<ol>
		<li>Clean the cantilevers at 80 &#176;C for 10 min by chromic acid treatment.</li>
		<li>Functionalize the cantilever by amino-silanization with 1% (v/v) APTES toluene solution and then bake the cantilever at 80 &#176;C for 15 min before conjugating to sulfo-SMCC.</li>
		<li>Link the C-ELP50nm-NGL to the surface with the maleimide group of sulfo-SMCC for 1.5 h.</li>
		<li>Wash away the unreacted C-ELP50nm-NGL on the coverslip by ultrapure water.</li>
		<li>Immerse a functionalized cantilever in 200 &#956;L of 50 &#956;M GL-CBM-XDoc protein solution containing 200 nM OaAEP1 at 25 &#176;C for 20-30 min. Then use AFM buffer (100 mM Tris, 100 mM NaCl, pH 7.4) to wash away unreacted protein. 
		NOTE: The surface chemistry of the cantilevers and the coverslip are similar.</li>
	</ol>
	</li>
</ol>
3. Stepwise polyprotein preparation with controlled sequences
<ol>
	<li>Link the ligation unit Coh-tev-L-POI-NGL to the GL-ELP50nm immobilized on the coverslip surface by OaAEP1 for 30 min.</li>
	<li>Use 15-20 mL of AFM buffer (100 mM Tris, 100 mM NaCl, pH 7.4) to wash away any unreacted proteins.</li>
	<li>Add 100 &#956;L of TEV protease (0.5 mM EDTA, 75 mM NaCl, 25 mM Tris-HCl 10% [v/v] glycerol, pH 8.0) to cleave the protein unit at the TEV recognize site for 1 h at 25 &#176;C.</li>
	<li>Use 15-20 mL of AFM buffer to wash away residual proteins.</li>
	<li>Link the ligation unit Coh-tev-L-POI-NGL to the GL-Ub-NGL-Glass by OaAEP1 for 30 min.</li>
	<li>Repeat steps 3.3 to 3.5 N-1 times to build protein construct GL-(Ub)n-NGL on the glass surface. Omit the last TEV cleavage reaction to reserve cohesin on the protein-polymer as Coh-tev-L-(Ub)n-NGL-Glass.</li>
</ol>
4. AFM Experiment measurement and data analysis
<ol>
	<li>AFM measurements
	<ol>
		<li>Add 1 mL of AFM buffer to the chamber with 10 mM CaCl2 and 5 mM Ascorbic Acid.</li>
		<li>Choose the D tip of the functionalized AFM probe for the experiment. Use the equipartition theorem to calibrate the cantilever in AFM buffer with an accurate spring constant (k) value before each experiment.</li>
		<li>Attach the cantilever tip to the sample surface to form the Cohesin/Dockerin pair.</li>
		<li>Retract the cantilever at a constant velocity of 400 nm&#183;s&#8722;1 from the surface. In the meantime, record the force-extension curve at a sample rate of 4000 Hz.</li>
	</ol>
	</li>
	<li>Data analysis
	<ol>
		<li>Use JPK data processing select force-extension traces.</li>
		<li>Use software to analyze the traces. Fit the curves with the worm-like-chain (WLC) model of polymer elasticity and obtain unfolding force and contour length increment for individual protein unfolding peak.</li>
		<li>Fit the histograms of unfolding forces with the Gaussian model to obtain the most probable values of unfolding force (&#60;Fu&#62;) and contour length increment (&#60;&#916;Lc&#62;).</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

We have described a protocol for enzymatic biosynthesis and immobilization of polyprotein and verified the polyprotein design by AFM-based SMFS. This methodology provides a novel approach to building the protein-polymer in a designed sequence, which complements previous methods for polyprotein engineering and immobilization4,6,52,53,54,<sup class="x...

Compared with synthetic polymers, natural multi-domain proteins have a uniform structure with a well-controlled number and type of subdomains1. This feature usually leads to improved biological function and stability2,3. Many approaches, such as cysteine-based disulfide bond coupling and recombinant DNA technology, have been developed for building such a polymerized protein with multiple domains4,5,6,7. However, the former method always results in a random and uncontrolled sequence, and the latter one leads to other problems, including the difficulty for the overexpression of toxic and large-size proteins and the purification of complex protein with cofactor and other delicate enzymes.
To meet this challenge, we develop an enzymatic method that conjugates protein monomer together for polymer/polyprotein in a stepwise fashion using a protein ligase OaAEP1 combined with a protease TEV8,9. OaAEP1 is a strict and efficient endopeptidase. Two proteins can be linked covalently as Asn-Gly-Leu sequence (NGL) through two termini by OaAEP1 in less than 30 min if the N-terminus is Gly-Leu residues(GL) and the other of which the C-terminus is NGL residues10. However, the use of OaAEP1 only to link protein monomer leads to a protein polymer with an uncontrolled sequence like the cysteine-based coupling method. Therefore, we design the N-terminus of the protein unit with a removable TEV protease site plus a leucine residue as ENLYFQ/G-L-POI. Before the TEV cleavage, the N-terminal would not participate in OaAEP1 ligation. And then the GL residues at N-terminus, which are compatible with further OaAEP1 ligation, is exposed after the TEV cleavage. Thus, we have achieved a sequential enzymatic biosynthesis method of polyprotein with a relatively well-controlled sequence.
Here, our stepwise enzymatic synthesis method can be used in polyprotein sample preparation, including sequence-controlled and uncontrolled, and protein immobilization for single-molecule studies as well, especially for the complex system such as metalloprotein.
Moreover, AFM-based SMFS experiments allow us to confirm the protein polymer construction and stability at the single-molecule level. Single-molecule force spectroscopy, including AFM, optical tweezer and magnetic tweezer, is a general tool in nanotechnology to manipulate biomolecule mechanically and measure their stability11,12,13,14,15,16,17,18,19,20. Single-molecule AFM has been widely used in the study of protein (un)folding21,22,23,24,25, the strength measurement of receptor-ligand interaction26,27,28,29,30,31,32,33,34,35, inorganic chemical bond20,36,37,38,39,40,41,42,43 and metal-ligand bond in metalloprotein44,45,46,47,48,49,50. Here, single-molecule AFM is used to verify the synthesized polyprotein sequence based on the corresponding protein unfolding signal.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>iron (III) chloride hexahydrate</td>
			<td>Energy chemical</td>
			<td></td>
			<td>99%</td>
		</tr>
		<tr>
			<td>Zinc chloride</td>
			<td>Alfa Aesar</td>
			<td></td>
			<td>100.00%</td>
		</tr>
		<tr>
			<td>calcium chloride hydrate</td>
			<td>Alfa Aesar</td>
			<td></td>
			<td>99.9965% crystalline aggregate</td>
		</tr>
		<tr>
			<td>L-Ascorbic Acid</td>
			<td>Sigma Life Science</td>
			<td></td>
			<td>Bio Xtra, &#8805;99.0%, crystalline</td>
		</tr>
		<tr>
			<td>(3-Aminopropyl) triethoxysilane</td>
			<td>Sigma-Aldrich</td>
			<td></td>
			<td>&#8805;99%</td>
		</tr>
		<tr>
			<td>sulfosuccinimidyl 4-(N-maleimidomethyl) cyclohexane-1-carboxylate</td>
			<td>Thermo Scientific</td>
			<td></td>
			<td>90%</td>
		</tr>
		<tr>
			<td>Glycerol</td>
			<td>Macklin</td>
			<td></td>
			<td>99%</td>
		</tr>
		<tr>
			<td>5,5&#39;-dithiobis(2-nitrobenzoic acid)</td>
			<td>Alfa Aesar</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Genes</td>
			<td>Genscript</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Nanowizard 4 AFM</td>
			<td>JPK Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>MLCT cantilever</td>
			<td>Bruker Corp</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Mono Q 5/50 GL</td>
			<td>GE Healthcare</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>AKTA FPLC system</td>
			<td>GE Healthcare</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glass coverslip</td>
			<td>Sail Brand</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Nanodrop 2000</td>
			<td>Thermo Scientific</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Avanti JXN-30 Centrifuge</td>
			<td>Beckman Coulter</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Gel Image System</td>
			<td>Tanon</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

The NGL residues introduced between adjacent proteins by OaAEP1 ligation will not affect protein monomer stability in the polymer as the unfolding force (&#60;Fu&#62;), and contour length increment (&#60;&#916;Lc&#62;) is comparable with the previous study (Figure 1). The purification result of the rubredoxin protein is shown in Figure 2. To prove the protein after TEV cleavage is compatible with the following OaAEP1 ligation to construct p...

Watch this Scientific Journal Video about OaAEP1-Mediated Enzymatic Synthesis and Immobilization of Polymerized Protein for Single-Molecule Force Spectroscopy at JoVE.com

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

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 ...

oaaep1-mediated-enzymatic-synthesis-immobilization-polymerized

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Biochemistry

6.6K Views.  Nanjing University. 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.

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

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. ...

Single-molecule Manipulation of G-quadruplexes by Magnetic Tweezers

Non-canonical nucleic acid secondary structure G-quadruplexes (G4) are involved in diverse cellular processes, such as DNA replication, transcription, RNA processing, and telomere elongation. During these processes, various proteins bind and resolve G4 structures to perform their function. As the function of G4 often depends on the stability of its folded structure, it is important to investigate how G4 binding proteins regulate the stability of G4. This work presents a method to manipulate single G4 molecules using magnetic tweezers, which enables studies of the regulation of G4 binding proteins on a single G4 molecule in real time. In general, this method is suitable for a wide scope of applications in studies for proteins/ligands interactions and regulations on various DNA or RNA secondary structures.

A single-molecule magnetic tweezers platform to manipulate G-quadruplexes is reported, which allows for the study of G4 stability and regulation by various proteins.

Non-canonical nucleic acid secondary structure G-quadruplexes (G4) are involved in diverse cellular processes, such as DNA replication, transcription, RNA ...

Using Three-color Single-molecule FRET to Study the Correlation of Protein Interactions

Single-molecule F&#246;rster resonance energy transfer (smFRET) has become a widely used biophysical technique to study the dynamics of biomolecules. For many molecular machines in a cell proteins have to act together&#160;with interaction partners in a functional cycle to fulfill their task. The extension of two-color to multi-color smFRET makes it possible to simultaneously probe more than one interaction or conformational change. This not only adds a new dimension to smFRET experiments but it also offers the unique possibility to directly study the sequence of events and to detect correlated interactions when using an immobilized sample and a total internal reflection fluorescence microscope (TIRFM). Therefore, multi-color smFRET is a versatile tool for studying biomolecular complexes in a quantitative manner and in a previously unachievable detail.
Here, we demonstrate how to overcome the special challenges of multi-color smFRET experiments on proteins. We present detailed protocols for obtaining the data and for extracting kinetic information. This includes trace selection criteria, state separation, and the recovery of state trajectories from the noisy data using a 3D ensemble Hidden Markov Model (HMM). Compared to other methods, the kinetic information is not recovered from dwell time histograms but directly from the HMM. The maximum likelihood framework allows us to critically evaluate the kinetic model and to provide meaningful uncertainties for the rates.
By applying our method to the heat shock protein 90 (Hsp90), we are able to disentangle the nucleotide binding and the global conformational changes of the protein. This allows us to directly observe the cooperativity between the two nucleotide binding pockets of the Hsp90 dimer.

Here, we present a protocol to obtain three-color smFRET data and its analysis with a 3D ensemble Hidden Markov Model. With this approach, scientists can extract kinetic information from complex protein systems, including cooperativity or correlated interactions.

Single-molecule F&#246;rster resonance energy transfer (smFRET) has become a widely used biophysical technique to study the dynamics of biomolecules. For ...

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 ...

A Fluorescence Fluctuation Spectroscopy Assay of Protein-Protein Interactions at Cell-Cell Contacts

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring cells. Of interest, only few assays are capable of specifically probing such interactions directly in living cells. Here, we present an assay to measure the binding of proteins expressed at the surfaces of neighboring cells, at cell-cell contacts. This assay consists of two steps: mixing of cells expressing the proteins of interest fused to different fluorescent proteins, followed by fluorescence fluctuation spectroscopy measurements at cell-cell contacts using a confocal laser scanning microscope. We demonstrate the feasibility of this assay in a biologically relevant context by measuring the interactions of the amyloid precursor-like protein 1 (APLP1) across cell-cell junctions. We provide detailed protocols on the data acquisition using fluorescence-based techniques (scanning fluorescence cross-correlation spectroscopy, cross-correlation number and brightness analysis) and the required instrument calibrations. Further, we discuss critical steps in the data analysis and how to identify and correct external, spurious signal variations, such as those due to photobleaching or cell movement.
In general, the presented assay is applicable to any homo- or heterotypic protein-protein interaction at cell-cell contacts, between cells of the same or different types and can be implemented on a commercial confocal laser scanning microscope. An important requirement is the stability of the system, which needs to be sufficient to probe diffusive dynamics of the proteins of interest over several minutes.

This protocol describes a fluorescence fluctuation spectroscopy-based approach to investigate interactions among proteins mediating cell-cell interactions, i.e. proteins localized in cell junctions, directly in living cells. We provide detailed guidelines on instrument calibration, data acquisition and analysis, including corrections to possible artefact sources.

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring ...

Proteome-wide Quantification of Labeling Homogeneity at the Single Molecule Level

Cell proteomes are often characterized using electrophoresis assays, where all species of proteins in the cells are non-specifically labeled with a fluorescent dye and are spotted by a photodetector following their separation. Single molecule fluorescence imaging can provide ultrasensitive protein detection with its ability for visualizing individual fluorescent molecules. However, the application of this powerful imaging method to electrophoresis assays is hampered by the lack of ways to characterize the homogeneity of fluorescent labeling of each protein species across the proteome. Here, we developed a method to evaluate the labeling homogeneity across the proteome based on a single molecule fluorescence imaging assay. In our measurement using a HeLa cell sample, the proportion of proteins labeled with at least one dye, which we termed &#8216;labeling occupancy&#8217; (LO), was determined to range from 50% to 90%, supporting the high potential of the application of single molecule imaging to sensitive and precise proteome analysis.

Here, we present a protocol to assess the labeling homogeneity for each protein species in a complex protein sample at the single molecule level.

Cell proteomes are often characterized using electrophoresis assays, where all species of proteins in the cells are non-specifically labeled with a ...

Single Molecule Fluorescence Energy Transfer Study of Ribosome Protein Synthesis

The ribosome is a large ribonucleoprotein complex that assembles proteins processively along mRNA templates. The diameter of the ribosome is approximately 20 nm to accommodate large tRNA substrates at the A-, P- and E-sites. Consequently, the ribosome dynamics are naturally de-phased quickly. Single molecule method can detect each ribosome separately and distinguish inhomogeneous populations, which is essential to reveal the complicated mechanisms of multi-component systems. We report the details of a smFRET method based on the Nikon Ti2 inverted microscope to probe the ribosome dynamics between the ribosomal protein L27 and tRNAs. The L27 is labeled at its unique Cys 53 position and reconstituted into a ribosome that is engineered to lack L27. The tRNA is labeled at its elbow region. As the tRNA moves to different locations inside the ribosome during the elongation cycle, such as pre- and post- translocation, the FRET efficiencies and dynamics exhibit differences, which have suggested multiple subpopulations. These subpopulations are not detectable by ensemble methods. The TIRF-based smFRET microscope is built on a manual or motorized inverted microscope, with home-built laser illumination. The ribosome samples are purified by ultracentrifugation, loaded into a home-built multi-channel sample cell and then illuminated via an evanescent laser field. The reflection laser spot can be used to achieve feedback control of perfect focus. The fluorescence signals are separated by a motorized filter-turret and collected by two digital CMOS cameras. The intensities are retrieved via the NIS-Elements software.

Single molecule fluorescence energy transfer is a method that tracks the tRNA dynamics during ribosomal protein synthesis. By tracking individual ribosomes, inhomogeneous populations are identified, which shed light on mechanisms. This method can be used to track biological conformational changes in general to reveal dynamic-function relationships in many other complexed biosystems. Single molecule methods can observe non-rate limiting steps and low-populated key intermediates, which are not accessible by conventional ensemble methods due to the average effect.

The ribosome is a large ribonucleoprotein complex that assembles proteins processively along mRNA templates. The diameter of the ribosome is approximately ...

An Optimized Single-Molecule Pull-Down Assay for Quantification of Protein Phosphorylation

Phosphorylation is a necessary posttranslational modification that regulates protein function and directs cell signaling outcomes. Current methods to measure protein phosphorylation cannot preserve the heterogeneity in phosphorylation across individual proteins. The single-molecule pull-down (SiMPull) assay was developed to investigate the composition of macromolecular complexes via immunoprecipitation of proteins on a glass coverslip followed by single-molecule imaging. The current technique is an adaptation of SiMPull that provides robust quantification of the phosphorylation state of full-length membrane receptors at the single-molecule level. Imaging thousands of individual receptors in this way allows for quantifying protein phosphorylation patterns. The present protocol details the optimized SiMPull procedure, from sample preparation to imaging. Optimization of glass preparation and antibody fixation protocols further enhances data quality. The current protocol provides code for the single-molecule data analysis that calculates the fraction of receptors phosphorylated within a sample. While this work focuses on phosphorylation of the epidermal growth factor receptor (EGFR), the protocol can be generalized to other membrane receptors and cytosolic signaling molecules.

The present protocol describes sample preparation and data analysis to quantify protein phosphorylation using an improved single-molecule pull-down (SiMPull) assay.

Phosphorylation is a necessary posttranslational modification that regulates protein function and directs cell signaling outcomes. Current methods to ...

Single-Molecule Diffusion and Assembly on Polymer-Crowded Lipid Membranes

Cellular membranes are highly crowded environments for biomolecular reactions and signaling. Yet, most in vitro experiments probing protein interaction with lipids employ naked bilayer membranes. Such systems lack the complexities of crowding by membrane-embedded proteins and glycans and exclude the associated volume effects encountered on cellular membrane surfaces. Also, the negatively charged glass surface onto which the lipid bilayers are formed prevents the free diffusion of transmembrane biomolecules. Here, we present a well-characterized polymer-lipid membrane as a mimic for crowded lipid membranes. This protocol utilizes polyethylene glycol (PEG)-conjugated lipids as a generalized approach for incorporating crowders into the supported lipid bilayer&#160;(SLB). First, a cleaning procedure of the microscopic slides and coverslips for performing single-molecule experiments is presented. Next, methods for characterizing the PEG-SLBs and performing single-molecule experiments of the binding, diffusion, and assembly of biomolecules using single-molecule tracking and photobleaching are discussed. Finally, this protocol demonstrates how to monitor the nanopore assembly of bacterial pore-forming toxin Cytolysin A (ClyA) on crowded lipid membranes with single-molecule photobleaching analysis. MATLAB codes with example datasets are also included to perform some of the common analyses such as particle tracking, extracting diffusive behavior, and subunit counting.

Here, a protocol to perform and analyze the binding, mobility, and assembly of single molecules on artificial crowded lipid membranes using single-molecule total internal reflection fluorescence (smTIRF) microscopy is presented.

Cellular membranes are highly crowded environments for biomolecular reactions and signaling. Yet, most in vitro experiments probing protein interaction ...

Use of Dual Optical Tweezers and Microfluidics for Single-Molecule Studies

Visual biochemistry is a powerful technique for observing the stochastic properties of single enzymes or enzyme complexes that are obscured in the averaging that takes place in bulk-phase studies. To achieve visualization, dual optical tweezers, where one trap is fixed and the other is mobile, are focused into one channel of a multi-stream microfluidic chamber positioned on the stage of an inverted fluorescence microscope. The optical tweezers trap single molecules of fluorescently labeled DNA and fluid flow through the chamber and past the trapped beads, stretches the DNA to B-form (under minimal force, i.e., 0 pN) with the nucleic acid being observed as a white string against a black background. DNA molecules are moved from one stream to the next, by translating the stage perpendicular to the flow to enable the initiation of reactions in a controlled manner. To achieve success, microfluidic devices with optically clear channels are mated to glass syringes held in place in a syringe pump. Optimal results use connectors permanently bonded to the flow cell, tubing that is mechanically rigid and chemically resistant, and which is connected to switching valves that eliminate bubbles that prohibit laminar flow.

Visual, single-molecule biochemistry studied through microfluidic chambers is greatly facilitated using glass barrel, gas-tight syringes, stable connections of tubing to flow cells, and elimination of bubbles by placing switching valves between the syringes and tubing. The protocol describes dual optical traps that enable visualization of DNA transactions and intermolecular interactions.