This work was supported by National Institutes of Health grant R15GM135904 (awarded to IM) and National Institutes of Health Grant GM110552 (awarded to DBO).

The authors have nothing to disclose.

Through the use of the FRET mapping methodology, we identified the signal sequence binding site on the SecA protein. Importantly, the presence of a 3-D crystal structure of the complex greatly facilitated our study. The strength of this mapping methodology lies in the ability to use an existing structure to identify locations for labeling. This methodology cannot be used to determine a 3-D structure; however, determination of structural elements56, refinement of an existing structure49, determination of a binding site location2,32, or elucidation of dynamic motion57, are all possible applications of this method.
In the SecYEG-SecA-PhoA complex, the three labeling sites form a triangle around the putative binding site (Figure 1). The employment of multiple distance measurements from the vertices of this triangle to the same residue refines the location information similar to GPS navigation methods. Three sites, SecA37, SecA341, and SecY292, were identified on SecA and SecY along with four sites, PhoA2, PhoA22, PhoA37, PhoA45 in the signal sequence and early mature region of the preprotein to give a total of 12 distances for mapping the location of the signal peptide (Table 1). Importantly, to improve the accuracy of the distance measurements, labeling sites should be located in relatively static regions of the protein, such as in secondary structure elements rather than loops. Furthermore, sites should also be in locations that are relatively accessible to solvent for ease and increased efficiency of labeling. Within the SecA-PhoA-SecYEG complex, the performance of distance measurements from the triangle sites or vertices to residues in the binding substrate were sufficient for locating the residues in the binding substrate to a relatively small area (Figure 3D,H and Figure 4D,H). Identification of the intersected area from the multiple distance measurements significantly refines the location from that of single distance measurement, as shown in Figure 3 and Figure 4. Thus, when using this method, the measurement of multiple distances is strongly recommended. Although this method can identify regions of molecules involved in binding, for example, it does not provide precise structural information; such information is best obtained from other structural methods such as X-ray, NMR, and cryo-EM. FRET distances can be used to effectively refine an existing structure18,19 or model, in this case, that was not possible as no model of the signal sequence bound to SecA exists.
To ensure that dye labeling occurs at only desired locations and FRET distance measurements are accurate, the use of mutagenesis for labeling is preferred. Generation of a Cys-less mutant requires relatively conservative mutation of Cys residues to Ser or similar residues in the otherwise wild-type protein using site-directed mutagenesis methods. In the current study, Cys mutations were introduced into Cys-free versions of SecA and SecYEG for labeling37. The activity of the mutant protein was verified with a growth assay followed by an in vitro malachite green ATPase assay41,42. Relevant activity assays depend on the function of the protein, for example for DNA binding proteins, a DNA binding assay would be appropriate58. Failure to ensure labeling at only one location can lead to labeling of more than one site with the same dye, which significantly complicates the distance determinations. Thus, the introduction of a second label on the same protein can be done through the incorporation of unnatural amino acid by site-directed mutagenesis. We employed this methodology to label the SecA-PhoA chimera at locations distinct from Cys residues by introducing the unnatural amino acid, p-azidophenylalanine and labeling with click chemistry39,40.
An additional important consideration of this method is the choice of dyes used and the associated R0 value. After identification of the potential labeling sites, the distances to be measured can be estimated from the 3-D structure. With this information, investigators can choose dyes pairs with R0 values that span the desired range of expected distances. For example, the AF488-AF647 dye pair has a calculated R0&#160;of 55.7 &#197; which provides a good range for labeling sites located an estimated 40-75 &#197; away from the putative binding site. Although the R0 values calculated in step 2.2 are useful for choosing which dye pair to use for your system, attachment of the dyes to the protein can alter their properties significantly. For greater accuracy, R0 values should be calculated from in situ experiments performed with labeled protein (steps 3.1 - 3.6.5).
Measurement of transfer efficiency can be done either by monitoring steady-state fluorescence emission and observing either a decrease in donor emission or an increase in acceptor emission.Although observation of both effects is desirable, the efficiency can be calculated from either as described5,8. Efficiency can also be calculated from the decrease in donor lifetime in the donor-acceptor sample relative to the donor-only sample. Determination of efficiency by more than one method is recommended, particularly the use of time-resolved methods to establish the relative homogeneity of the efficiencies measured.
The mapping method also allowed us to determine the relative orientation of the signal sequence and the early mature regions of the PhoA preprotein relative to SecA and the putative binding site. A SecA-SecYEG X-ray crystal structure and subsequent cryo-EM study provided clarity regarding the structure of the signal sequence and the early mature region of the preprotein with respect to the channel and SecA34,35. In the X-ray structure, residues 1-41 of the OmpA preprotein were attached to the tip of the two-helix finger and visualized in a hairpin structure in the channel (shown in pink, Figure 5). The locations of the four PhoA residues in the SecA-PhoA chimera were mapped onto this structure using the same protocol as described above. As shown in Figure 5, the location of the PhoA37 and PhoA45 residues (yellow, orange, red) is in between PhoA2 and PhoA22, with PhoA45 closer to PhoA2. These findings, particularly the location of PhoA45, suggested that the PhoA preprotein was forming a hairpin structure.
To further validate our FRET-identified binding site, we performed a comparison of our mapped locations with that of the OmpA preprotein, by excising the 41 residue preprotein structure from the channel and modeling it into the regions defined by FRET mapping (Figure 5, cyan). Without any alteration of the preprotein X-ray structure, we find that the locations of residues 2, 22, and 37 (shown in blue, green, and yellow) on the OmpA preprotein fragment excised structure agree remarkably well with the FRET mapped locations (Figure 5B) and suggest that the hairpin forms prior to channel entry. The OmpA preprotein ends at residue 41 in the X-ray crystal structure; however, the C-terminal of SecY, which is unstructured, provides an indication of the possible location of PhoA45. In our modeled structure, the hairpin loop sits at the mouth of the channel, poised to facilitate the translocation of the preprotein across the membrane. Thus, in this example, the FRET mapping methodology enhances existing information of the static SecA-SecYEG structure, by providing a hint of the dynamic motion needed for protein translocation across the membrane. Although not suitable for de novo structure determinations, if 3-dimensional structural information is available, the FRET mapping methodology can further the current understanding of structure-function relationships, through elucidation of binding sites and dynamic motions.

For proteins, elucidation of dynamics along with 3-dimensional (3-D) structural knowledge leads to an enhanced understanding of structure-function relationships of biomolecular systems. Structural methods, such as X-ray crystallography and cryogenic electron microscopy, capture a static structure and often require the determination of multiple structures to elucidate aspects of biomolecule binding and dynamics1. This article discusses a solution-based method for mapping global structural elements, such as binding sites or binding interactions, that are potentially more transient and less easily captured by static methods. Strong candidate systems for this methodology are ones in which a 3-D structure has been previously determined by X-ray crystallography, NMR spectroscopy, or other structural methods. In this case, we take advantage of the X-ray crystal structure of the SecA-SecYEG complex, a central player in the protein general secretory pathway, to map the location of a signal peptide binding site using F&#246;rster resonance energy transfer (FRET) prior to the transport of the preprotein across the membrane2. Manipulation of the biological system through genetic modifications coupled with our knowledge of the 3-D structure enabled the determination of the conformation of the signal sequence and early mature region immediately prior to insertion into the channel 3.
FRET involves the radiation-less transfer of energy from one molecule (donor) to another (acceptor) in a distance-dependent fashion that is through space4,5. The efficiency of this transfer is monitored through either a decrease in donor or an increase in acceptor fluorescence intensity. The efficiency of energy transfer can be described as
E = R06/(R06 + R6)
in which the R0 value is the distance at which the transfer is 50% efficient6. The technique has previously been described as a molecular ruler and is effective at determining distances in the 2.5-12 nm range, depending on the identity of the donor-acceptor dyes4,7,8,9. The donor fluorescence intensities and lifetimes with or without acceptor allow determination of transfer efficiencies and consequently, distances5,8. Due to the availability of the technology, sensitivity of the method, and ease of use, FRET has also found broad application in such areas as single-molecule fluorescence spectroscopy and confocal microscopy6. The advent of fluorescent proteins such as green fluorescent protein has made the observation of intracellular dynamics and live-cell imaging relatively facile10,11. Many FRET applications such as these are discussed in detail in this virtual issue.
In this study, we particularly focus on the use of FRET measurements to yield distance values to determine structural details. Previously, FRET measurements have been effectively used to determine the conformation of DNA molecules when bound to protein12,13,14, the internal dynamics of proteins, and protein binding interactions15,16,17. The advantages of this method lie in the ability to determine flexible and dynamic structural elements in a solution with relatively low amounts of material. Significantly, this method is particularly effective when used in conjunction with existing structural information and cannot be used as a means of 3-D structure determination. The method provides the best insight and refinement of structure if the work builds on existing structural information often coupled with computational simulation18,19. Here, the use of distances obtained from steady-state and time-resolved FRET measurements is described to map a binding site, the location of which was not known, on an existing crystallographic structure of the SecA-SecYEG complex, major proteins in the general secretory pathway3.
The general secretory pathway, a highly conserved system from prokaryotes to eukaryotes to archaea, mediates the transport of proteins either across or into the membrane to their functional location in the cell. For Gram-negative bacteria, such as E. coli, the organism used in our study, proteins are inserted into or translocated across the inner membrane to the periplasm. The bacterial SecY channel complex (termed the translocon) coordinates with other proteins to translocate the newly synthesized protein, which is directed to its correct location in the cell through a signal sequence typically located at the N-terminus20,21. For proteins bound for the periplasm, the ATPase SecA protein associates with the exit tunnel of the ribosome, and with the preprotein after approximately 100 residues have been translated22. Along with the SecB chaperone protein, it maintains the preprotein in an unfolded state. SecA binds to the SecYEG translocon, and through many cycles of ATP hydrolysis, facilitates protein transport across the membrane23,24.
SecA is a multi-domain protein that exists in cytosolic and membrane-bound forms. A homodimeric protein in the cytosol, SecA consists of a preprotein binding or cross-linking domain25, two nucleotide-binding domains, a helical wing domain, a helical scaffold domain, and the two helix finger (THF)26,27,28,29 (Figure 1). In previous crystallographic studies of the SecA-SecYEG complex, the location of the THF suggested that it was actively involved in protein translocation and subsequent cross-linking experiments with the signal peptide further established the significance of this region in protein translocation30,31. Previous studies, using the FRET mapping methodology, demonstrated that exogenous signal peptides bind to this region of SecA2,32. To fully understand the conformation and location of the signal sequence and early mature region of the preprotein prior to insertion into the SecYEG channel, a protein chimera in which the signal sequence and residues of the early mature region were attached to SecA through a Ser-Gly linker was created (Figure 1). Using this biologically viable construct, it was further demonstrated that the signal sequence and early mature region of the preprotein bind to the THF in a parallel fashion2. Subsequently, the FRET mapping methodology was used to elucidate the conformation and location of the signal sequence and early mature region in the presence of SecYEG as described below3.
Knowledge of the 3-D structure of the SecA-SecYEG complex33,34,35 and the possible location of the binding site allowed us to judiciously place donor-acceptor labels in locations where the intersection of individual FRET distances identifies the binding site location. These FRET mapping measurements revealed that the signal sequence and the early mature region of the preprotein form a hairpin with the tip located at the mouth of the SecYEG channel, demonstrating that the hairpin structure is templated prior to channel insertion.

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			<td>490 nm LED laser</td>
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			<td>Alexa Fluor 647 C2 Maleimide//DIBO Alkyne</td>
			<td>Life Technologies</td>
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			<td>Alexa Fluor 488 DIBO Alkyne</td>
			<td>Life Technologies</td>
			<td>S10904</td>
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			<td>S10906</td>
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			<td>GlobalsWE</td>
			<td>Laboratory for Fluorescence Dynamics, University of California, Irvine</td>
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			<td>spectral analysis program for time-resolved decays</td>
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			<td>H&#173;4&#173;Azido&#173;Phe&#173;OH</td>
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f&#246;rster resonance energy transfer mapping: a new methodology to elucidate global structural features

F&#246;rster resonance energy transfer (FRET) is an established fluorescence-based method used to successfully measure distances in and between biomolecules in vitro as well as within cells. In FRET, the efficiency of energy transfer, measured by changes in fluorescence intensity or lifetime, relates to the distance between two fluorescent molecules or labels. Determination of dynamics and conformational changes from the distances are just some examples of applications of this method to biological systems. Under certain conditions, this methodology can add to and enhance existing X-ray crystal structures by providing information regarding dynamics, flexibility, and adaptation to binding surfaces. We describe the use of FRET and associated distance determinations to elucidate structural properties, through the identification of a binding site or the orientations of dimer subunits. Through judicious choice of labeling sites, and often employment of multiple labeling strategies, we have successfully applied these mapping methods to determine global structural properties in a protein-DNA complex and the SecA-SecYEG protein translocation system. In the SecA-SecYEG system, we have used FRET mapping methods to identify the preprotein-binding site and determine the local conformation of the bound signal sequence region. This study outlines the steps for performing FRET mapping studies, including identification of appropriate labeling sites, discussion of possible labels including non-native amino acid residues, labeling procedures, how to perform measurements, and interpreting the data.

Watch this Scientific Journal Video about FÃ¶rster Resonance Energy Transfer Mapping: A New Methodology to Elucidate Global Structural Features at JoVE.com

F&#246;rster Resonance Energy Transfer Mapping: A New Methodology to Elucidate Global Structural Features

The study details the methodology of FRET mapping including the selection of labeling sites, choice of dyes, acquisition, and data analysis. This methodology is effective at determining binding sites, conformational changes, and dynamic motions in protein systems and is most useful if performed in conjunction with existing 3-D structural information.

F&#246;rster resonance energy transfer (FRET) is an established fluorescence-based method used to successfully measure distances in and between ...

frster-resonance-energy-transfer-mapping-new-methodology-to-elucidate

Nanyang Technological University

Research

JoVE Journal

Biochemistry

2.4K Views.  Wesleyan University. The study details the methodology of FRET mapping including the selection of labeling sites, choice of dyes, acquisition, and data analysis. This methodology is effective at determining binding sites, conformational changes, and dynamic motions in protein systems and is most useful if performed in conjunction with existing 3-D structural information.

Video: Förster Resonance Energy Transfer Mapping: A New Methodology to Elucidate Global Structural Features

Protein Complex Affinity Capture from Cryomilled Mammalian Cells

Affinity capture is an effective technique for isolating endogenous protein complexes for further study. When used in conjunction with an antibody, this technique is also frequently referred to as immunoprecipitation. Affinity capture can be applied in a bench-scale and in a high-throughput context. When coupled with protein mass spectrometry, affinity capture has proven to be a workhorse of interactome analysis. Although there are potentially many ways to execute the numerous steps involved, the following protocols implement our favored methods. Two features are distinctive: the use of cryomilled cell powder to produce cell extracts, and antibody-coupled paramagnetic beads as the affinity medium. In many cases, we have obtained superior results to those obtained with more conventional affinity capture practices. Cryomilling avoids numerous problems associated with other forms of cell breakage. It provides efficient breakage of the material, while avoiding denaturation issues associated with heating or foaming. It retains the native protein concentration up to the point of extraction, mitigating macromolecular dissociation. It reduces the time extracted proteins spend in solution, limiting deleterious enzymatic activities, and it may reduce the non-specific adsorption of proteins by the affinity medium. Micron-scale magnetic affinity media have become more commonplace over the last several years, increasingly replacing the traditional agarose- and Sepharose-based media. Primary benefits of magnetic media include typically lower non-specific protein adsorption; no size exclusion limit because protein complex binding occurs on the bead surface rather than within pores; and ease of manipulation and handling using magnets.

Here we describe protocols to disrupt mammalian cells by solid-state milling at a cryogenic temperature, produce a cell extract from the resulting cell powder, and isolate protein complexes of interest by affinity capture upon antibody-coupled micron-scale paramagnetic beads.

Affinity capture is an effective technique for isolating endogenous protein complexes for further study. When used in conjunction with an antibody, this ...

Atomic Scale Structural Studies of Macromolecular Assemblies by Solid-state Nuclear Magnetic Resonance Spectroscopy

Supramolecular protein assemblies play fundamental roles in biological processes ranging from host-pathogen interaction, viral infection to the propagation of neurodegenerative disorders. Such assemblies consist in multiple protein subunits organized in a non-covalent way to form large macromolecular objects that can execute a variety of cellular functions or cause detrimental consequences. Atomic insights into the assembly mechanisms and the functioning of those macromolecular assemblies remain often scarce since their inherent insolubility and non-crystallinity often drastically reduces the quality of the data obtained from most techniques used in structural biology, such as X-ray crystallography and solution Nuclear Magnetic Resonance (NMR). We here present magic-angle spinning solid-state NMR spectroscopy (SSNMR) as a powerful method to investigate structures of macromolecular assemblies at atomic resolution. SSNMR can reveal atomic details on the assembled complex without size and solubility limitations. The protocol presented here describes the essential steps from the production of 13C/15N isotope-labeled macromolecular protein assemblies to the acquisition of standard SSNMR spectra and their analysis and interpretation. As an example, we show the pipeline of a SSNMR structural analysis of a filamentous protein assembly.

Structures of supramolecular protein assemblies at atomic resolution are of high relevance because of their crucial roles in a variety of biological phenomena. Herein, we present a protocol to perform high-resolution structural studies on insoluble and non-crystalline macromolecular protein assemblies by magic-angle spinning solid-state nuclear magnetic resonance spectroscopy (MAS SSNMR).

Supramolecular protein assemblies play fundamental roles in biological processes ranging from host-pathogen interaction, viral infection to the ...

Electrochemical Detection of Deuterium Kinetic Isotope Effect on Extracellular Electron Transport in Shewanella oneidensis MR-1

Direct electrochemical detection of c-type cytochrome complexes embedded in the bacterial outer membrane (outer membrane c-type cytochrome complexes; OM c-Cyts) has recently emerged as a novel whole-cell analytical method to characterize the bacterial electron transport from the respiratory chain to the cell exterior, referred to as the extracellular electron transport (EET). While the pathway and kinetics of the electron flow during the EET reaction have been investigated, a whole-cell electrochemical method to examine the impact of cation transport associated with EET has not yet been established. In the present study, an example of a biochemical technique to examine the deuterium kinetic isotope effect (KIE) on EET through OM c-Cyts using a model microbe, Shewanella oneidensis MR-1, is described. The KIE on the EET process can be obtained if the EET through OM c-Cyts acts as the rate-limiting step in the microbial current production. To that end, before the addition of D2O, the supernatant solution was replaced with fresh media containing a sufficient amount of the electron donor to support the rate of upstream metabolic reactions, and to remove the planktonic cells from a uniform monolayer biofilm on the working electrode. Alternative methods to confirm the rate-limiting step in microbial current production as EET through OM c-Cyts are also described. Our technique of a whole-cell electrochemical assay for investigating proton transport kinetics can be applied to other electroactive microbial strains.

Electrochemical Detection of Deuterium Kinetic Isotope Effect on Extracellular Electron Transport in Shewanella oneidensis MR-1

Here we present a protocol of whole-cell electrochemical experiments to study the contribution of proton transport to the rate of extracellular electron transport via the outer-membrane cytochromes complex in Shewanella oneidensis MR-1.

Direct electrochemical detection of c-type cytochrome complexes embedded in the bacterial outer membrane (outer membrane c-type cytochrome complexes; OM ...

Using In Vitro Fluorescence Resonance Energy Transfer to Study the Dynamics Of Protein Complexes at a Millisecond Time Scale

Proteins are the primary operators of biological systems, and they usually interact with other macro- or small molecules to carry out their biological functions. Such interactions can be highly dynamic, meaning the interacting subunits are constantly associated and dissociated at certain rates. While measuring the binding affinity using techniques such as quantitative pull-down reveals the strength of the interaction, studying the binding kinetics provides insights on how fast the interaction occurs and how long each complex can exist. Furthermore, measuring the kinetics of an interaction in the presence of an additional factor, such as a protein exchange factor or a drug, helps reveal the mechanism by which the interaction is regulated by the other factor, providing important knowledge for the advancement of biological and medical research. Here, we describe a protocol for measuring the binding kinetics of a protein complex that has a high intrinsic association rate and can be dissociated quickly by another protein. The method uses fluorescence resonance energy transfer to report the formation of the protein complex in vitro, and it enables monitoring the fast association and dissociation of the complex in real time on a stopped-flow fluorimeter. Using this assay, the association and dissociation rate constants of the protein complex are quantified.

Protein-protein interactions are critical for biological systems, and studies of the binding kinetics provide insights into the dynamics and function of protein complexes. We describe a method that quantifies the kinetic parameters of a protein complex using fluorescence resonance energy transfer and the stopped-flow technique. &#160;

Proteins are the primary operators of biological systems, and they usually interact with other macro- or small molecules to carry out their biological ...

Removal and Replacement of Endogenous Ligands from Lipid-Bound Proteins and Allergens

Many major allergens bind to hydrophobic lipid-like molecules, including Mus m 1, Bet v 1, Der p 2, and Fel d 1. These ligands are strongly retained and have the potential to influence the sensitization process either through directly stimulating the immune system or altering the biophysical properties of the allergenic protein. In order to control for these variables, techniques are required for the removal of endogenously bound ligands and, if necessary, replacement with lipids of known composition. The cockroach allergen Bla g 1 encloses a large hydrophobic cavity which binds a heterogeneous mixture of endogenous lipids when purified using traditional techniques. Here, we describe a method through which these lipids are removed using reverse-phase HPLC followed by thermal annealing to yield Bla g 1 in either its Apo-form or reloaded with a user-defined mixture of fatty acid or phospholipid cargoes. Coupling this protocol with biochemical assays reveal that fatty acid cargoes significantly alter the thermostability and proteolytic resistance of Bla g 1, with downstream implications for the rate of T-cell epitope generation and allergenicity. These results highlight the importance of lipid removal/reloading protocols such as the one described herein when studying allergens from both recombinant and natural sources. The protocol is generalizable to other allergen families including lipocalins (Mus m 1), PR-10 (Bet v 1), MD-2 (Der p 2) and Uteroglobin (Fel d 1), providing a valuable tool to study the role of lipids in the allergic response.

This protocol describes the removal of endogenous lipids from allergens, and their replacement with user-specified ligands through reverse-phase HPLC coupled with thermal annealing. 31P-NMR and circular dichroism allow for the rapid confirmation of ligand removal/loading, and the recovery of native allergen structure.

Many major allergens bind to hydrophobic lipid-like molecules, including Mus m 1, Bet v 1, Der p 2, and Fel d 1. These ligands are strongly retained and ...

NMR-Based Fragment Screening in a Minimum Sample but Maximum Automation Mode

Fragment-based screening (FBS) is a well-validated and accepted concept within the drug discovery process both in academia and industry. The greatest advantage of NMR-based fragment screening is its ability not only to detect binders over 7-8 orders of magnitude of affinity but also to monitor purity and chemical quality of the fragments and thus to produce high quality hits and minimal false positives or false negatives. A prerequisite within the FBS is to perform initial and periodic quality control of the fragment library, determining solubility and chemical integrity of the fragments in relevant buffers, and establishing multiple libraries to cover diverse scaffolds to accommodate various macromolecule target classes (proteins/RNA/DNA). Further, an extensive NMR-based screening protocol optimization with respect to sample quantities, speed of acquisition and analysis at the level of biological construct/fragment-space, in condition-space (buffer, additives, ions, pH, and temperature) and in ligand-space (ligand analogues, ligand concentration) is required. At least in academia, these screening efforts have so far been undertaken manually in a very limited fashion, leading to limited availability of screening infrastructure not only in the drug development process but also in the context of chemical probe development. In order to meet the requirements economically, advanced workflows are presented. They take advantage of the latest state-of-the-art advanced hardware, with which the liquid sample collection can be filled in a temperature-controlled fashion into the NMR-tubes in an automated manner. 1H/19F NMR ligand-based spectra are then collected at a given temperature. High-throughput sample changer (HT sample changer) can handle more than 500 samples in temperature-controlled blocks. This together with advanced software tools speeds up data acquisition and analysis. Further, application of screening routines on protein and RNA samples are described to make aware of the established protocols for a broad user base in biomacromolecular research.

Fragment-based screening by NMR is a robust method to rapidly identify small molecule binders to biomacromolecules (DNA, RNA, or proteins). Protocols describing automation-based sample preparation, NMR experiments &#38; acquisition conditions, and analysis workflows are presented. The technique allows for optimal exploitation of both 1H and 19F NMR-active nuclei for detection.

Fragment-based screening (FBS) is a well-validated and accepted concept within the drug discovery process both in academia and industry. The greatest ...

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 ...

Pure Shift Nuclear Magnetic Resonance: a New Tool for Plant Metabolomics

Nuclear Magnetic Resonance (NMR) is one of the most powerful tools used in metabolomics. It stands as a highly accurate and reproducible method that not only provides quantitative data but also permits structural identification of the metabolites present in complex mixtures.
Metabolic profiling by 1H NMR has proven useful in the study of various types of plant scenarios, which include the evaluation of crop&#160;conditions, harvest and post- harvest treatments, metabolic phenotyping, metabolic pathways, gene regulation, identification of biomarkers, chemotaxonomy, quality control, denomination of origin, among others. However, signal overlapping of the large number of resonances with expanded J-coupling multiplicities complicates the spectra analysis and its interpretation, and represents a limitation for classical 1H NMR profiling.
In the last decade, novel NMR broadband homonuclear decoupling techniques through which multiplet signals collapse into single resonance lines - commonly called Pure Shift methods - have been developed to overcome the spectra resolution problem inherent to 1H NMR classical spectra.
Here a step-by-step protocol of the plant extract preparation and the procedure to record optimal Pure Shift PSYCHE and SAPPHIRE-PSYCHE spectra in three different plant matrices - Vanilla plant&#160;leaves, potato tubers (S. tuberosum), and Cape gooseberries (P. peruviana) - is presented. The effect of the gain in resolution in metabolic identification, correlation analysis and multivariate analyses, as compared against classical spectra, is discussed.

This paper presents the use of PSYCHE and SAPPHIRE-PSYCHE in the metabolic profiling of plants and includes detailed procedures for sample preparation and optimal Pure Shift NMR spectra recording. Examples through which the gain in resolution achieved by homonuclear decoupling allows a more comprehensive understanding of the system are discussed.

Nuclear Magnetic Resonance (NMR) is one of the most powerful tools used in metabolomics. It stands as a highly accurate and reproducible method that not ...

F&#246;rster Resonance Energy Transfer Measurements in Living Plant Cells

Sensitized emission-based F&#246;rster resonance energy transfer (FRET) experiments are easily done but depend on the microscopic setup. Confocal laser scanning microscopes have become a workhorse for biologists. Commercial systems offer high flexibility in laser power adjustment and detector sensitivity and often combine different detectors to obtain the perfect image. However, the comparison of intensity-based data from different experiments and setups is often impossible due to this flexibility. Biologist-friendly procedures are of advantage and allow for simple and reliable adjustment of laser and detector settings. 
Furthermore, as FRET experiments in living cells are affected by the variability in protein expression and donor-acceptor ratios, protein expression levels must be considered for data evaluation. Described here is a simple protocol for reliable and reproducible FRET measurements, including routines for the estimation of protein expression and adjustment of laser intensity and detector settings. Data evaluation will be performed by calibration with a fluorophore fusion of known FRET efficiency. To improve simplicity, correction factors have been compared that have been obtained in cells and by measuring recombinant fluorescent proteins.

A protocol is provided for setting up a standard confocal laser-scanning microscope for in vivo F&#246;rster resonance energy transfer measurements, followed by data evaluation.

Sensitized emission-based F&#246;rster resonance energy transfer (FRET) experiments are easily done but depend on the microscopic setup. Confocal laser ...

Time-resolved F&#246;rster Resonance Energy Transfer Assays for Measurement of Endogenous Phosphorylated STAT Proteins in Human Cells

The Janus kinase (JAK)/signal transducer and activator of transcription (STAT) signaling pathway plays a crucial role in mediating cellular responses to cytokines and growth factors. STAT proteins are activated by tyrosine phosphorylation mediated mainly by JAKs. The abnormal activation of STAT signaling pathways is implicated in many human diseases, especially cancer and immune-related conditions. Therefore, the ability to monitor STAT protein phosphorylation within the native cell signaling environment is important for both academic and drug discovery research. The traditional assay formats available to quantify phosphorylated STAT proteins include western blotting and the enzyme-linked immunosorbent assay (ELISA). These heterogeneous methods are labor-intensive, low-throughput, and often not reliable (specific) in the case of western blotting. Homogeneous (no-wash) methods are available but remain expensive. 
Here, detailed protocols are provided for the sensitive, robust, and cost-effective measurement in a 384-well format of endogenous levels of phosphorylated STAT1 (Y701), STAT3 (Y705), STAT4 (Y693), STAT5 (Y694/Y699), and STAT6 (Y641) in cell lysates from adherent or suspension cells using the novel THUNDER time-resolved F&#246;rster resonance energy transfer (TR-FRET) platform. The workflow for the cellular assay is simple, fast, and designed for high-throughput screening (HTS). The assay protocol is flexible, uses a low-volume sample (15 &#181;L), requires only one reagent addition step, and can be adapted to low-throughput and high-throughput applications. Each phospho-STAT sandwich immunoassay is validated under optimized conditions with known agonists and inhibitors and generates the expected pharmacology and Z'-factor values. As TR-FRET assays are ratiometric and require no washing steps, they provide much better reproducibility than traditional approaches. Together, this suite of assays provides new cost-effective tools for a more comprehensive analysis of specific phosphorylated STAT proteins following cell treatment and the screening and characterization of specific and selective modulators of the JAK/STAT signaling pathway.

Time-resolved F&#246;rster resonance energy transfer cell-based assay protocols are described for the simple, specific, sensitive, and robust quantification of endogenous phosphorylated signal transducer and activator of transcription (STAT) 1/3/4/5/6 proteins in cell lysates in a 384-well format.

The Janus kinase (JAK)/signal transducer and activator of transcription (STAT) signaling pathway plays a crucial role in mediating cellular responses to ...

Förster Resonance Energy Transfer Mapping: A New Methodology to Elucidate Global Structural Features

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Protein Complex Affinity Capture from Cryomilled Mammalian Cells

Atomic Scale Structural Studies of Macromolecular Assemblies by Solid-state Nuclear Magnetic Resonance Spectroscopy

Electrochemical Detection of Deuterium Kinetic Isotope Effect on Extracellular Electron Transport in Shewanella oneidensis MR-1

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Single Molecule Fluorescence Energy Transfer Study of Ribosome Protein Synthesis

Pure Shift Nuclear Magnetic Resonance: a New Tool for Plant Metabolomics

Förster Resonance Energy Transfer Measurements in Living Plant Cells

Time-resolved Förster Resonance Energy Transfer Assays for Measurement of Endogenous Phosphorylated STAT Proteins in Human Cells