We thank members of the Cuevas-Velazquez lab for the critical review of the manuscript. This work was supported by Programa de Apoyo a Proyectos de Investigaci&#243;n e Innovaci&#243;n Tecnol&#243;gica, Direcci&#243;n General de Asuntos del Personal Acad&#233;mico, Universidad Nacional Aut&#243;noma de M&#233;xico (UNAM-PAPIIT) project number IA203422; Consejo Nacional de Humanidades, Ciencias y Tecnolog&#237;a (CONAHCYT), project number 252952; and Programa de Apoyo a la Investigaci&#243;n y el Posgrado, Facultad de Qu&#237;mica, Universidad Nacional Aut&#243;noma de M&#233;xico, Grant 5000-9182. CET (CVU 1083636) and CAPD (CVU 1269643) acknowledge CONAHCYT for their M.Sc. Scholarship.

1. Plasmid construct
<ol>
	<li>Amplify the open reading frame (ORF) that codes for the desired IDR by polymerase chain reaction (PCR). Do not include the stop codon since the ORF will be flanked by the genes that encode the fluorescent proteins. For the amplification, design primers with the SacI (5&#39;) and BglII (3&#39;) restriction sites. 
	NOTE: For the representative results section, we used AtLEA4-5 as the selected IDR. We amplified the ORF of AtLEA4-5 from the pTrc99A-AtLEA4-5 plasmid12.</li>
	<li>Digest the ORF PCR product by consecutive restriction with SacI and BglII enzymes13.</li>
	<li>Obtain the commercial plasmid pDRFLIP38-AtLEA4-5 (#178189) from Addgene (https://www.addgene.org/178189/).</li>
	<li>Conduct a consecutive restriction using SacI and BglII enzymes to remove the AtLEA4-5 ORF from the pDRFLIP38-AtLEA4-5 plasmid, leaving an open plasmid containing the FRET pair13.</li>
	<li>Purify the digested DNA fragments using gel recovery from an agarose gel electrophoresis14. Ligate the restricted fragments using DNA ligase15.</li>
	<li>Transform the cloning reaction into competent Escherichia coli cells (DH5&#945; or related strains) and select in Luria-Bertani (LB) plates containing 50 &#181;g/mL ampicillin16. Grow overnight (ON) at 37 &#176;C.</li>
	<li>Isolate and grow at least five transformed colonies in a new plate and genotype by PCR17. Purify the plasmid DNA from a positive transformed colony using standard miniprep methods18.</li>
	<li>Verify the extraction and integrity of the plasmid DNA using agarose gel electrophoresis19. Verify the correct cloning of the construct using Sanger sequencing20.</li>
</ol>
2. Plasmid expression in yeast cells
NOTE: Use standard aseptic techniques to perform the following steps. Use a culture laminar flow hood or a lighter.
<ol>
	<li>Inoculate a streak of S. cerevisiae BJ5465 (ATCC: 208289) strain into 3 mL of Yeast Peptone Dextrose (YPD) medium and grow ON&#160;at 30 &#176;C and 200 rpm. 
	NOTE: BJ5465 is a protease-deficient (CH1) strain recommended for working with IDRs. Other S. cerevisiae strains can be tested and used.</li>
	<li>Centrifuge 1 mL of the overnight saturated (OD600 ~ 3-4) yeast culture at 14,000 x g for 1 min. Carefully remove the supernatant by pipetting.</li>
	<li>Resuspend the pellet in 1 mL of Tris-EDTA (TE) buffer (pH 7.5) by gently flicking the tube. Centrifuge at 14,000 x g for 1 min and carefully remove the supernatant.</li>
	<li>Resuspend the pellet in 500 &#181;L of Lazy Bones buffer (40% w/v PEG 3,350; 100&#160;mM lithium acetate; 10 mM Tris-HCl; 1 mM EDTA; pH 7.5) by pipetting. Add 25 &#181;L of boiled (100 &#176;C for 5 min, followed by cooling on ice for 5 min) salmon sperm DNA (2 mg/mL) to the resuspended yeast cells.</li>
	<li>Add 100 ng of the plasmid DNA to the mixture and vortex the mixture for 1 min to ensure thorough mixing. Allow the mixture to stand at room temperature for 1-2 h.</li>
	<li>Heat shock the yeast mixture at 42 &#176;C for 12 min, then cool on ice for another 12 min. Centrifuge the tube at 14 000 x g for 1 min and carefully remove the supernatant.</li>
	<li>Resuspend the pellet in 1 mL of TE buffer (pH 7.5). Plate 100 &#181;L of the transformed cells onto Yeast Synthetic-dropout Medium without uracil (SD-ura) plates supplemented with 15 g/L agar.</li>
	<li>Incubate plates at 30 &#176;C for 2-3 days (Figure 1). Streak at least five yeast transformants in a new plate and grow ON.</li>
</ol>
3. Validation of yeast transformants
<ol>
	<li>Pick a small portion of each transformant candidate by gently scraping and resuspending in 5 &#181;L of 20&#160;mM NaOH into individual PCR tubes.</li>
	<li>Heat the sample at 99 &#176;C for 10 min in a thermal cycler to lyse the cells and release the DNA into the solution. This step is crucial for preparing the DNA template for PCR.</li>
	<li>Transfer 1 &#181;L of the boiled sample to a separate PCR reaction mix containing the appropriate primers and PCR reagents for genotyping. We used the following primers: Forward primer: 5&#180;-AAATATACCCCAGCCTCGATCTAGA-3&#180;. Reverse primer: 5&#180;-GTAATACGACTCACTATAGGGCG-3&#180;.</li>
	<li>Set up a PCR reaction and validate the presence of the correct plasmid using agarose gel electrophoresis. Select one transformant for further experiments.</li>
</ol>
4. Preparation of yeast cell culture for FRET assay
<ol>
	<li>Inoculate a streak of the selected yeast transformant into 3 mL of liquid 1x SD-Ura medium. 
	NOTE: Do this procedure in a laminar flow hood and sterile material.</li>
	<li>Grow at 30 &#176;C, 200 rpm for at least 12 h to reach saturation. Measure the OD600. Growth should fall between OD600 = 1.0-2.0. 
	&#8203;NOTE: If OD600 is lower than 1.0, give cells more incubation time to grow. If OD600 exceeds 2.0, do not proceed and restart from step 4.1.</li>
</ol>
5. Yeast cell preparation for FRET assay
<ol>
	<li>Collect 2 mL of the overnight yeast culture and centrifuge at 14,000 x g at room temperature. Carefully remove the supernatant by pipetting.</li>
	<li>Resuspend the pellet in 1 mL of 50 mM 2-(N-Morpholino) ethanesulfonic acid (MES) buffer (pH 6).</li>
	<li>Centrifuge at 14,000 x g at room temperature. Remove the supernatant.</li>
	<li>Repeat steps 5.2 and 5.3. Resuspend yeast cells in 2 mL of MES buffer, pH 6 and pour the volume into a reagent reservoir.</li>
</ol>
6. Setting up the fluorescence measurements
NOTE: The present scanning is considered to be performed in a microplate reader with fluorescence mode.
<ol>
	<li>For basic settings, set the instrument as follows: Measurement type: Fluorescence Intensity (FI) spectrum; Microplate name: Greiner 96 F-bottom.</li>
	<li>For optic settings, set the instrument as follows: No. of wavelength scan points: 91; Excitation wavelength: 433 nm; Excitation bandwidth: 10 nm; Emission wavelength: 460 - 550; Step width: 1 nm; Emission bandwidth: 10 nm; Focal height obtained by autofocus; Focal height: 5 mm; Wavelength used for gain: 490 nm.</li>
	<li>For general settings, set the instrument as follows: Settling time: 0.1 s; Reading direction: unidirectional, horizontal left to right, top to bottom. 
	&#8203;NOTE: The manually entered gain must be adjusted for every measurement using the well expected to display the highest fluorescence intensity levels throughout the wavelength scan.</li>
</ol>
7. Preparation of hypertonic solutions and fluorescence measurements
<ol>
	<li>Prepare solutions of 0 M, 0.2 M, 0.4 M, 0.6 M, 0.8 M, 1 M, 1.5 M, and 2 M NaCl in a 96-well clear-bottom black plate. The final volume in each well will be 200 &#181;L (150 &#181;L of osmolyte solution + 50 &#181;L of yeast cell suspension). 
	NOTE: Other osmolytes can be used to induce hyperosmotic stress. All the solutions must be prepared in 50 mM MES buffer pH 6.</li>
	<li>Load 150 &#181;L of 50 mM MES buffer, pH 6 in first three wells of row A (A1, A2, A3) and in the subsequent wells, add 150 &#181;L of the corresponding solution in increasing order from left to right (Figure 2). To test all the concentrations suggested, continue the loading in row B. 
	NOTE: This setting considers the measurements of 3 technical replicates for each concentration.</li>
	<li>Use a 12-channel micropipette to transfer 50 &#181;L of washed yeast cells to each well. Yeast cells tend to sediment to the bottom of the reservoir. Make sure to resuspend the yeast suspension thoroughly by pipetting up and down before transferring the cells.</li>
	<li>Load cells into the 96-well plate containing the different solutions and mix well by pipetting up and down at least 4x.</li>
	<li>Measure fluorescence intensity emission spectra immediately for the desired wells in a microplate reader with the specification settings of step 6. Repeat the procedure for every IDR to be tested in the FRET assay.</li>
	<li>Optional step. If available, measure a donor-only construct (pDRFLIP38-mCerulean3) following the steps described here. 
	NOTE: We recommended performing this step to calculate each condition&#39;s FRET efficiency. If the reader cannot perform this step, FRET can be calculated using the FRET ratio method. Both methods for FRET calculations are explained in steps 8 and 9.</li>
	<li>Perform at least three replicates on three independent days for every construct.</li>
</ol>
8. Data processing using the FRET efficiency method
NOTE: For readers with knowledge of the R programming language, we provide a set of R scripts to perform data processing described in this section. Scripts can be found at&#160;https://github.com/Kaz-bits/cuevaslab-procotols/tree/main/FRET. Follow the instructions in the README file.
<ol>
	<li>Export data as .xlsx&#160;files from the microplate reader.</li>
	<li>Normalize fluorescence intensity values at every scan wavelength to the isosbestic point of the FRET pair used. 
	NOTE: This step is necessary for comparing the spectra of all conditions tested. The isosbestic point of mCerulean3 and Citrine FRET pair is 515 nm. For example, obtain the ratio of fluorescence intensity values of 460 nm/515 nm, 461 nm/515 nm, 462 nm/515 nm, and so on.</li>
	<li>Calculate the mean for every technical replicate once the fluorescence values are normalized. In total, there should be a column for every condition of hyperosmotic stress.</li>
	<li>Visualize all the fluorescence intensity spectra in the same plot (Figure 3). Each spectrum should display a combination of the fluorescence emission spectra of individual mCerulean3 and Citrine. In rare cases, where the FRET efficiency is 0, only the donor fluorescence emission spectrum will be observed, or when FRET efficiency is 1, only the acceptor fluorescence will be observed. The fluorescence emission peak for mCerulean3 (donor) is 475 nm, and the fluorescence emission peak for Citrine (acceptor) is 525 nm.</li>
	<li>Determine if the fluorescence measurement displays a typical FRET change in response to hyperosmotic stress. If the conformation of the IDR is sensitive to the treatment, this will reflect as a change in FRET efficiency. A typical change in FRET efficiency will couple a decrease in the fluorescence intensity of the donor with an increase in the fluorescence intensity of the acceptor or vice versa.</li>
	<li>Quantify the FRET efficiency (EFRET). Obtain the donor (mCerulean3) normalized peak values (475 nm/515 nm) for the IDR construct and the donor-only construct for every condition of hyperosmotic stress. Use the mean normalized peak values of the technical replicates. Perform this operation for each of the three independent replicates. Calculate EFRET with the following formula: 
	<img alt="Equation 1" src="/files/ftp_upload/66275/66275eq01.jpg" style="margin: auto;" /> 
	Where IDA is the ratio 475 nm/515 nm from the IDR construct (FRET pair construct), and ID is the ratio 475 nm/515 nm from the donor-only construct.</li>
	<li>Compare FRET efficiencies. Normalize the EFRET values of every hyperosmotic stress condition to the EFRET of the non-stress condition (for example, 0 M NaCl). Make comparisons using a box plot or a smooth curve plot. Perform a One-way ANOVA followed by a post hoc Tukey statistical test (p-values: *p &#60; 0.05, **p &#60; 0.01, ***p &#60; 0.001, ****p &#60; 0.0001).</li>
</ol>
9. Data processing using the FRET ratio method
NOTE: If donor-only measurements cannot be acquired, perform the FRET ratio method. This method compares the FRET ratio across the different hyperosmotic stress conditions. Steps 7.1 to 7.5 should be performed before the steps of this section to validate a typical FRET behavior. For readers with knowledge of the R programming language, we provide a set of R scripts to perform data processing described in this section. Scripts can be found at&#160;https://github.com/Kaz-bits/cuevaslab-procotols/tree/main/FRET. Follow the instructions in the README file.
<ol>
	<li>Extract data at 475 nm and 525 nm from the xlsx&#160;file previously exported from the microplate reader. These values correspond to the fluorescence emission peak for mCerulean3 (donor, DxDm) and the fluorescence emission peak for Citrine (acceptor, DxAm) when the donor fluorophore is excited (433 nm).</li>
	<li>Normalize fluorescence intensity values at 475 nm and 525 nm to the isosbestic point of the FRET pair used. The isosbestic point of mCerulean3 and Citrine FRET pair is 515 nm. Obtain the FRET ratio (DxAm/DxDm).</li>
	<li>Compare FRET ratios. Normalize the DxAm/DxDm values of every hyperosmotic stress condition to the mean DxAm/DxDm of the non-stress condition (for example, 0 M NaCl). Make comparisons using a box plot or a smooth curve plot (Figure 4 and Figure 5). Perform a One-way ANOVA followed by a post hoc Tukey statistical test (p-values: *p &#60; 0.05, **p &#60; 0.01, ***p &#60; 0.001, ****p &#60; 0.0001).</li>
</ol>

FRET Analysis of IDR Structural Sensitivity to Study Hyperosmotic Stress in Yeast

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The authors declare no conflicts of interest.

The method presented here offers a way to obtain insights into how the global dimensions of the ensemble of IDRs sense and respond to environmental perturbations. This method relies on a genetically encoded construct and requires no additional components beyond a plasmid stable expression in yeast cells, making it adaptable for potential applications in other cell types. Moreover, it is versatile for exploring other physicochemical perturbations that eukaryotic cells experience during their life cycle21...

Intrinsically disordered regions (IDRs) are critical components in cellular processes1. In combination with structured domains, IDRs are essential for protein functions. The amino acid composition of IDRs is biased, represented mainly by charged, hydrophilic, and small residues. Because of this property, IDRs are considered low complexity domains2,3. Numerous IDRs have garnered attention, primarily because these regions play a crucial role in pathological conditions, particularly neurodegenerative diseases. Such diseases are characterized by self-assembly and subsequent extracellular or intracellular deposition of IDRs in neurons4. Examples of such IDRs include amyloid-&#946;&#160;(A&#946;) in Alzheimer&#39;s disease, huntingtin (HTT) in Huntington&#39;s disease, and TAR DNA-binding protein-43 (TDP-43) and fused in sarcoma (FUS) in amyotrophic lateral sclerosis and frontotemporal dementia4. The study of the structural rearrangements of IDRs in the context of disease has been significantly enhanced by spectroscopic methods, including fluorescence resonance energy transfer (FRET).
The hydrophilic and extended nature of IDRs makes them extremely sensitive to changes in the physicochemical properties of the solution environment5. The degree by which the conformational ensemble of IDRs is modified by the environment is called structural sensitivity5,6,7. Different techniques can be used to study the conformation and dynamics of IDRs, including circular dichroism (CD) and small-angle X-ray scattering (SAXS)8,9. Unfortunately, CD and SAXS require large quantities of purified proteins, so they are not appropriate for studies in cells. In contrast, FRET is a technique that measures the fluorescence intensity of two fluorescent molecules that specifically label one IDR, meaning that they can be monitored in complex mixtures such as living cells10. Dynamically measuring the structural sensitivity of IDRs in living cells is necessary for understanding how the environment regulates the conformation and function of the disordered proteome.
FRET is a powerful method for quantifying the structural sensitivity of IDRs, as well as globular and multidomain proteins in living cells. The method requires a construct consisting of an IDR of interest sandwiched between two fluorescent proteins (FP), known as a FRET pair. For this protocol, we suggest the use of mCerulean3 as the donor FP and Citrine as the acceptor FP, because of their large dynamic range, compared to other FPs reported in a previous study about IDRs sensitivity6. FRET has previously been exploited to measure the structural sensitivity of a plant IDR in different cellular contexts6. In addition, this technique has been used to characterize overall protein dimensions of IDRs by different research groups both in vitro and in vivo5,11.
Here, we describe the ensemble FRET method for studying the structural sensitivity of IDRs in living yeast (Saccharomyces cerevisiae) cells. We show representative results that are based in a plant IDR called AtLEA4-5. AtLEA4-5 is disordered in solution, but folds into &#945;-helix when macromolecular crowding is induced in vitro12. AtLEA4-5 is a good reference model for this method because it is relatively small (158 residues), disordered and sensitive to environmental perturbation as reported in silico and in vitro6,12. The method presented here can be scaled for high throughput approaches because yeast cells are easy to grow, and the treatment is applied in small volumes. In addition, small modifications to the protocol can be applied to other cellular systems such as bacteria and plant cells6. The protocol can be performed in any molecular biology laboratory with access to a microplate reader with fluorescence mode, an equipment available in most research institutions.

<table><tbody><tr><td>96-well plate</td><td>Greiner Bio-One</td><td>655096</td><td></td></tr><tr><td>Agar</td><td>Sigma-Aldrich</td><td>5040</td><td></td></tr><tr><td>BglII</td><td>New England BioLabs</td><td>R0144S</td><td></td></tr><tr><td>BJ5465 cells</td><td>American Type Culture Collection</td><td>208289</td><td></td></tr><tr><td>Buffer MES 50 mM</td><td>Sigma-Aldrich</td><td>M8250</td><td></td></tr><tr><td>Buffer Tris-HCl 10 mM</td><td>Invitrogen</td><td>15506017</td><td></td></tr><tr><td>EDTA 1 mM</td><td>Merck</td><td>108452</td><td></td></tr><tr><td>Falcon tubes</td><td>Corning</td><td>352057</td><td></td></tr><tr><td>LB media</td><td>Sigma-Aldrich</td><td>L2897</td><td></td></tr><tr><td>Lithium acetate 0.1 M</td><td>Sigma-Aldrich</td><td>L6883</td><td></td></tr><tr><td>Low Melt Agarose</td><td>GOLDBIO</td><td>A-204-25</td><td></td></tr><tr><td>Microcentrifuge</td><td>eppendorf</td><td>5452000010</td><td></td></tr><tr><td>Miniprep kit</td><td>ZymoPure</td><td>D4210</td><td></td></tr><tr><td>NaOH 0.02 M</td><td>Merck</td><td>106462</td><td></td></tr><tr><td>PEG 3,350 40%</td><td>Sigma-Aldrich</td><td>1546547</td><td></td></tr><tr><td>plasmid pDRFLIP38-AtLEA4-5</td><td>addgene</td><td>178189</td><td></td></tr><tr><td>Plate reader</td><td>BMG LABTECH</td><td>CLARIOstar Plus</td><td></td></tr><tr><td>SacI</td><td>New England BioLabs</td><td>R3156S</td><td></td></tr><tr><td>Salmon sperm DNA 2 mg/mL</td><td>Thermo Fisher Scientific</td><td>15632011</td><td></td></tr><tr><td>SD-Ura</td><td>Sigma-Aldrich</td><td>Y1501</td><td></td></tr><tr><td>Sodium cloride</td><td>Sigma-Aldrich</td><td>S9888</td><td></td></tr><tr><td>Taq polymesare</td><td>Promega</td><td>M5123</td><td></td></tr><tr><td>Transiluminator</td><td>Accuris instruments</td><td>E4000</td><td></td></tr><tr><td>UV-Visible spectrophotometer</td><td>Thermo Fisher Scientific</td><td>Biomate3</td><td></td></tr><tr><td>YPD media</td><td>Sigma-Aldrich</td><td>Y1500</td><td></td></tr></tbody></table>

estimation of structural sensitivity of intrinsically disordered regions in response to hyperosmotic stress in living cells using fret

Intrinsically disordered regions (IDRs) are protein domains that participate in crucial cellular processes. During stress conditions, the physicochemical properties of the cellular environment change, directly impacting the conformational ensemble of IDRs. IDRs are inherently sensitive to environmental perturbations. Studying how the physicochemical properties of the cell regulate the conformational ensemble of IDRs is essential for understanding the environmental control of their function. Here, we describe a step-by-step method for measuring the structural sensitivity of IDRs in living Saccharomyces cerevisiae cells in response to hyperosmotic stress conditions. We present the use of ensemble fluorescence resonance energy transfer (FRET) to estimate how the global dimensions of IDRs change during a progressive increase of hyperosmotic stress imposed on cells with any osmolyte. In addition, we provide a script for processing fluorescence measurements and comparing structural sensitivity for different IDRs. By following this method, researchers can obtain valuable insights into the conformational changes that IDRs undergo in the complex intracellular milieu upon changing environments.

After transforming yeast cells with pDRFLIP38-AtLEA4-5 plasmid, the fluorescence of the positive transformants was observed with a blue light transilluminator and a filter (Figure 1). Preparing the different solutions to induce hyperosmotic stress is time-consuming, so we suggest following the 96-well template of Figure 2. Immediately after the hyperosmotic stress treatment with varying concentrations of sodium chloride, the fluorescence emission spectra were ac...

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Author Spotlight: Unlocking the World of Intrinsically Disordered Regions with Cellular Sensing and Responses

Intrinsically disordered regions (IDRs) are flexible protein domains that modify their conformation in response to environmental changes. Ensemble fluorescence resonance energy transfer (FRET) can estimate protein dimensions under different conditions. We present a FRET approach to assess IDR structural sensitivity in living Saccharomyces cerevisiae cells under hyperosmotic stress.

Estimation of Structural Sensitivity of Intrinsically Disordered Regions in Response to Hyperosmotic Stress in Living Cells Using FRET

Intrinsically disordered regions (IDRs) are protein domains that participate in crucial cellular processes. During stress conditions, the physicochemical ...

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Research

JoVE Journal

Biochemistry

920 Views.  Universidad Nacional Autónoma de México. We streven ernaar te begrijpen hoe intrinsiek ongeordende regio's veranderingen in de fysisch-chemische eigenschappen van de omgeving waarnemen en erop reageren. We combineren biofysische, biochemische, genetische en celbiologische technieken om te monitoren hoe het structurele ensemble van ongeordende regio's verandert in levende cellen. Eiwitbiofysische technieken worden momenteel gebruikt om de bevestiging van IVR's te bestuderen.Deze omvatten nucleaire magnetische resonantie, circu...

Video: Auteur in de schijnwerpers: de wereld van intrinsiek ongeordende regio's ontsluiten met cellulaire detectie en reacties

Structural Information from Single-molecule FRET Experiments Using the Fast Nano-positioning System

Single-molecule F&#246;rster Resonance Energy Transfer (smFRET) can be used to obtain structural information on biomolecular complexes in real-time. Thereby, multiple smFRET measurements are used to localize an unknown dye position inside a protein complex by means of trilateration. In order to obtain quantitative information, the Nano-Positioning System (NPS) uses probabilistic data analysis to combine structural information from X-ray crystallography with single-molecule fluorescence data to calculate not only the most probable position but the complete three-dimensional probability distribution, termed posterior, which indicates the experimental uncertainty. The concept was generalized for the analysis of smFRET networks containing numerous dye molecules. The latest version of NPS, Fast-NPS, features a new algorithm using Bayesian parameter estimation based on Markov Chain Monte Carlo sampling and parallel tempering that allows for the analysis of large smFRET networks in a comparably short time. Moreover, Fast-NPS allows the calculation of the posterior by choosing one of five different models for each dye, that account for the different spatial and orientational behavior exhibited by the dye molecules due to their local environment.
Here we present a detailed protocol for obtaining smFRET data and applying the Fast-NPS. We provide detailed instructions for the acquisition of the three input parameters of Fast-NPS: the smFRET values, as well as the quantum yield and anisotropy of the dye molecules. Recently, the NPS has been used to elucidate the architecture of an archaeal open promotor complex. This data is used to demonstrate the influence of the five different dye models on the posterior distribution.

We present the setup and experimental procedure to obtain smFRET data from large donor-acceptor networks with a TIRF microscope. The step-by-step analysis of these measurements with the Bayesian inference software Fast-NPS yields high-resolved structural information via the application of adapted dye models.

Single-molecule F&#246;rster Resonance Energy Transfer (smFRET) can be used to obtain structural information on biomolecular complexes in real-time. ...

High Precision FRET at Single-molecule Level for Biomolecule Structure Determination

A protocol on how to perform high-precision interdye distance measurements using F&#246;rster resonance energy transfer (FRET) at the single-molecule level in multiparameter fluorescence detection (MFD) mode is presented here. MFD maximizes the usage of all &#34;dimensions&#34; of fluorescence to reduce photophysical and experimental artifacts and allows for the measurement of interdye distance with an accuracy up to ~1 &#197; in rigid biomolecules. This method was used to identify three conformational states of the ligand-binding domain of the N-methyl-D-aspartate (NMDA) receptor to explain the activation of the receptor upon ligand binding. When comparing the known crystallographic structures with experimental measurements, they agreed within less than 3 &#197; for more dynamic biomolecules. Gathering a set of distance restraints that covers the entire dimensionality of the biomolecules would make it possible to provide a structural model of dynamic biomolecules.

A protocol for high-precision FRET experiments at the single molecule level is presented here. Additionally, this methodology can be used to identify three conformational states in the ligand-binding domain of the N-methyl-D-aspartate (NMDA) receptor. Determining precise distances is the first step towards building structural models based on FRET experiments.

A protocol on how to perform high-precision interdye distance measurements using F&#246;rster resonance energy transfer (FRET) at the single-molecule ...

Visualizing the Conformational Dynamics of Membrane Receptors Using Single-Molecule FRET

The ability of cells to respond to external signals is essential for cellular development, growth, and survival. To respond to a signal from the environment, a cell must be able to recognize and process it. This task mainly relies on the function of membrane receptors, whose role is to convert signals into the biochemical language of the cell. G protein-coupled receptors (GPCRs) constitute the largest family of membrane receptor proteins in humans. Among GPCRs, metabotropic glutamate receptors (mGluRs) are a unique subclass that function as obligate dimers and possess a large extracellular domain that contains the ligand-binding site. Recent advances in structural studies of mGluRs have improved the understanding of their activation process. However, the propagation of large-scale conformational changes through mGluRs during activation and modulation is poorly understood. Single-molecule fluorescence resonance energy transfer (smFRET) is a powerful technique to visualize and quantify the structural dynamics of biomolecules at the single-protein level. To visualize the dynamic process of mGluR2 activation, fluorescent conformational sensors based on unnatural amino acid (UAA) incorporation were developed that allowed site-specific protein labeling without perturbation of the native structure of receptors. The protocol described here explains how to perform these experiments, including the novel UAA labeling approach, sample preparation, and smFRET data acquisition and analysis. These strategies are generalizable and can be extended to investigate the conformational dynamics of a variety of membrane proteins.

This study presents a detailed procedure to perform single-molecule fluorescence resonance energy transfer (smFRET) experiments on G protein-coupled receptors (GPCRs) using site-specific labeling via unnatural amino acid (UAA) incorporation. The protocol provides a step-by-step guide for smFRET sample preparation, experiments, and data analysis.

The ability of cells to respond to external signals is essential for cellular development, growth, and survival. To respond to a signal from the ...

Measurement of Force-Sensitive Protein Dynamics in Living Cells Using a Combination of Fluorescent Techniques

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called mechanotransduction. A crucial step in mechanotransduction is the transmission of forces between the external and internal environments. To transmit forces, there must be a sustained, unbroken physical linkage created by a series of protein-protein interactions. For a given protein-protein interaction, mechanical load can either have no effect on the interaction, lead to faster disassociation of the interaction, or even stabilize the interaction. Understanding how molecular load dictates protein turnover in living cells can provide valuable information about the mechanical state of a protein, in turn elucidating its role in mechanotransduction. Existing techniques for measuring force-sensitive protein dynamics either lack direct measurements of protein load or rely on the measurements performed outside of the cellular context. Here, we describe a protocol for the F&#246;rster resonance energy transfer-fluorescence recovery after photobleaching (FRET-FRAP) technique, which enables the measurement of force-sensitive protein dynamics within living cells. This technique is potentially applicable to any FRET-based tension sensor, facilitating the study of force-sensitive protein dynamics in variety of subcellular structures and in different cell types.

Here, we present a protocol for the simultaneous use of F&#246;rster resonance energy transfer-based tension sensors to measure protein load and fluorescence recovery after photobleaching to measure protein dynamics enabling the measurement of force-sensitive protein dynamics within living cells.

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called ...

Bioengineering

Visualizing Intracellular SNARE Trafficking by Fluorescence Lifetime Imaging Microscopy

Soluble N-ethylmaleimide sensitive fusion protein (NSF) attachment protein receptor (SNARE) proteins are key for membrane trafficking, as they catalyze membrane fusion within eukaryotic cells. The SNARE protein family consists of about 36 different members. Specific intracellular transport routes are catalyzed by specific sets of 3 or 4 SNARE proteins that thereby contribute to the specificity and fidelity of membrane trafficking. However, studying the precise function of SNARE proteins is technically challenging, because SNAREs are highly abundant and functionally redundant, with most SNAREs having multiple and overlapping functions. In this protocol, a new method for the visualization of SNARE complex formation in live cells is described. This method is based on expressing SNARE proteins C-terminally fused to fluorescent proteins and measuring their interaction by F&#246;rster resonance energy transfer (FRET) employing fluorescence lifetime imaging microscopy (FLIM). By fitting the fluorescence lifetime histograms with a multicomponent decay model, FRET-FLIM allows (semi-)quantitative estimation of the fraction of the SNARE complex formation at different vesicles. This protocol has been successfully applied to visualize SNARE complex formation at the plasma membrane and at endosomal compartments in mammalian cell lines and primary immune cells, and can be readily extended to study SNARE functions at other organelles in animal, plant, and fungal cells.

This protocol describes a new method allowing for the quantitative visualization of complex formation of SNARE proteins, based on F&#246;rster resonance energy transfer, and fluorescence lifetime imaging microscopy.

Soluble N-ethylmaleimide sensitive fusion protein (NSF) attachment protein receptor (SNARE) proteins are key for membrane trafficking, as they catalyze ...

Immunology and Infection

Automated Two-dimensional Spatiotemporal Analysis of Mobile Single-molecule FRET Probes

Single-molecule F&#246;rster resonance energy transfer (smFRET) is a versatile technique reporting on distances in the sub-nanometer to nanometer range. It has been used in a wide range of biophysical and molecular biological experiments, including the measurement of molecular forces, characterization of conformational dynamics of biomolecules, observation of intracellular colocalization of proteins, and determination of receptor-ligand interaction times. In a widefield microscopy configuration, experiments are typically performed using surface-immobilized probes. Here, a method combining single-molecule tracking with alternating excitation (ALEX) smFRET experiments is presented, permitting the acquisition of smFRET time traces of surface-bound, yet mobile probes in plasma membranes or glass-supported lipid bilayers. For the analysis of recorded data, an automated, open-source software collection was developed supporting (i) the localization of fluorescent signals, (ii) single-particle tracking, (iii) determination of FRET-related quantities including correction factors, (iv) stringent verification of smFRET traces, and (v) intuitive presentation of the results. The generated data can conveniently be used as input for further exploration via specialized software, e.g., for the assessment of the diffusional behavior of probes or the investigation of FRET transitions.

This article presents a method for spatiotemporal analysis of mobile, single-molecule F&#246;rster resonance energy transfer (smFRET)-based probes using widefield fluorescence microscopy. The newly developed software toolkit allows the determination of smFRET time traces of moving probes, including the correct FRET efficiency and the molecular positions, as functions of time.

Single-molecule F&#246;rster resonance energy transfer (smFRET) is a versatile technique reporting on distances in the sub-nanometer to nanometer range. ...

Coupled Assays for Monitoring Protein Refolding in Saccharomyces cerevisiae

Proteostasis, defined as the combined processes of protein folding/biogenesis, refolding/repair, and degradation, is a delicate cellular balance that must be maintained to avoid deleterious consequences 1. External or internal factors that disrupt this balance can lead to protein aggregation, toxicity and cell death. In humans this is a major contributing factor to the symptoms associated with neurodegenerative disorders such as Huntington's, Parkinson's, and Alzheimer's diseases 10. It is therefore essential that the proteins involved in maintenance of proteostasis be identified in order to develop treatments for these debilitating diseases. This article describes techniques for monitoring in vivo protein folding at near-real time resolution using the model protein firefly luciferase fused to green fluorescent protein (FFL-GFP). FFL-GFP is a unique model chimeric protein as the FFL moiety is extremely sensitive to stress-induced misfolding and aggregation, which inactivates the enzyme 12. Luciferase activity is monitored using an enzymatic assay, and the GFP moiety provides a method of visualizing soluble or aggregated FFL using automated microscopy. These coupled methods incorporate two parallel and technically independent approaches to analyze both refolding and functional reactivation of an enzyme after stress. Activity recovery can be directly correlated with kinetics of disaggregation and re-solubilization to better understand how protein quality control factors such as protein chaperones collaborate to perform these functions. In addition, gene deletions or mutations can be used to test contributions of specific proteins or protein subunits to this process. In this article we examine the contributions of the protein disaggregase Hsp104 13, known to partner with the Hsp40/70/nucleotide exchange factor (NEF) refolding system 5, to protein refolding to validate this approach.

Coupled Assays for Monitoring Protein Refolding in Saccharomyces cerevisiae

This article describes the use of a firefly luciferase-GFP fusion protein to investigate in vivo protein folding in Saccharomyces cerevisiae. Using this reagent, refolding of a model heat-denatured protein can be monitored simultaneously by fluorescence microscopy and an enzymatic assay to probe the roles of proteostasis network components in protein quality control.

Proteostasis, defined as the combined processes of protein folding/biogenesis, refolding/repair, and degradation, is a delicate cellular balance that must ...

Biology

Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing molecules of interest to fluorescent proteins such as GFP to study their expression, localization and function. However, GFP and its derivatives are significantly larger and less photostable than organic fluorophores generally used for in vitro experiments, and this can limit the scope of investigation. 
We recently introduced a straightforward, versatile and high-throughput method based on electroporation, allowing the internalization of biomolecules labeled with organic fluorophores into living microorganisms. Here we describe how to use electroporation to internalize labeled DNA fragments or proteins into Escherichia coli and Saccharomyces cerevisi&#230;, how to quantify the number of internalized molecules using fluorescence microscopy, and how to quantify the viability of electroporated cells. Data can be acquired at the single-cell or single-molecule level using fluorescence or FRET. The possibility of internalizing non-labeled molecules that trigger a physiological observable response in vivo is also presented. Finally, strategies of optimization of the protocol for specific biological systems are discussed.

Studies of biomolecules in vivo are crucial for understanding molecular function in a biological context. Here we describe a novel method allowing the internalization of fluorescent biomolecules, such as DNA or proteins, into living microorganisms. Analysis of in vivo data recorded by fluorescence microscopy is also presented and discussed.

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing ...

FRET Imaging in Three-dimensional Hydrogels

Imaging of F&#246;rster resonance energy transfer (FRET)&#160;is a powerful tool for examining cell biology in real-time. Studies utilizing FRET commonly employ two-dimensional (2D) culture, which does not mimic the three-dimensional (3D) cellular microenvironment. A method to perform quenched emission FRET imaging using conventional widefield epifluorescence microscopy of cells within a 3D hydrogel environment is presented. Here an analysis method for ratiometric FRET probes that yields linear ratios over the probe activation range is described. Measurement of intracellular cyclic adenosine monophosphate (cAMP) levels is demonstrated in chondrocytes under forskolin stimulation using a probe for EPAC1 activation (ICUE1) and the ability to detect differences in cAMP signaling dependent on hydrogel material type, herein a photocrosslinking hydrogel (PC-gel, polyethylene glycol dimethacrylate) and a thermoresponsive hydrogel (TR-gel). Compared with 2D FRET methods, this method requires little additional work. Laboratories already utilizing FRET imaging in 2D can easily adopt this method to perform cellular studies in a 3D microenvironment. It can further be applied to high throughput drug screening in engineered 3D microtissues. Additionally, it is compatible with other forms of FRET imaging, such as anisotropy measurement and fluorescence lifetime imaging (FLIM), and with advanced microscopy platforms using confocal, pulsed, or modulated illumination.

F&#246;rster resonance energy transfer (FRET) imaging is a powerful tool for real-time cell biology studies. Here a method for FRET imaging cells in physiologic three-dimensional (3D) hydrogel microenvironments using conventional epifluorescence microscopy is presented. An analysis for ratiometric FRET probes that yields linear ratios over the activation range is described.

Imaging of F&#246;rster resonance energy transfer (FRET)&#160;is a powerful tool for examining cell biology in real-time. Studies utilizing FRET commonly ...

Detecting and Characterizing Protein Self-Assembly In Vivo by Flow Cytometry

Protein self-assembly governs protein function and compartmentalizes cellular processes in space and time. Current methods to study it suffer from low-sensitivity, indirect read-outs, limited throughput, and/or population-level rather than single-cell resolution. We designed a flow cytometry-based single methodology that addresses all of these limitations: Distributed Amphifluoric FRET or DAmFRET. DAmFRET detects and quantifies protein self-assemblies by sensitized emission FRET in vivo, enables deployment across model systems-from yeast to human cells-and achieves sensitive, single-cell, high-throughput read-outs irrespective of protein localization or solubility.

This article describes a FRET-based flow cytometry protocol to quantify protein self-assembly in both S. cerevisiae and HEK293T cells.

Estimation of Structural Sensitivity of Intrinsically Disordered Regions in Response to Hyperosmotic Stress in Living Cells Using FRET

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