The work in V.C.&#39;s laboratory is supported by grants from NIH (R35GM144059 and R01GM50224), NSF (MCB1913165 and IOS1758046), and BARD (IS-5276-20) to V.C.

Nicotiana benthamiana, Agrobacterium tumefaciens strain EHA105, or GV3101 were used for the present study.
1. FRET vector construction
<ol>
	<li>Select fluorescent tags for the donor/acceptor FRET pair.
	<ol>
		<li>Use EGFP from pPZP-RCS2A-DEST-EGFP-N115,28 (see Table of Materials) to generate the donor vector.</li>
		<li>Use mRFP from pPZP-RCS2A-DEST-mRFP-N1 (see Table of Materials) to generate the acceptor vector.</li>
	</ol>
	</li>
	<li>Generate the donor/acceptor FRET constructs using a site-specific recombination cloning technique34, such as the Gateway recombination cloning system35.
	<ol>
		<li>Amplify the coding sequences of the proteins of interest36 (i.e., the Arabidopsis OTLD1 and LSH10)15. 
		NOTE: It is also a good idea to utilize a negative control that represents a homolog of one of the interacting proteins but is not expected to exhibit interaction; the OTLD1-LSH10 interaction study employs a homolog of LSH10,&#160;LSH4, that does not recognize OTLD1. OTLD1, LSH10, and LSH4 cDNAs are amplified by PCR using primers listed in Table 1.</li>
		<li>Clone OTLD1, LSH10, and LSH4 into the entry vector pDONR207 by the site-specific recombination cloning technique34.</li>
		<li>Use the Gateway LR Clonase II (see Table of Materials) to transfer LSH10 and LSH4 from pDONR207 into the binary destination vector pPZP-RCS2A-DEST-EGFP-N1 to generate the binary donor constructs p35S::LSH10-GFP (tested construct) and p35S::LSH4-GFP (negative control).</li>
		<li>Use the same commercially available enzyme (step 1.2.3) to transfer OTLD1 from pDONR207 into the binary destination vector pPZP-RCS2A-DEST-mRFP-N1 to generate the binary acceptor construct p35S::OTLD1-mRFP (tested construct).</li>
		<li>PCR-amplify36 mRFP from pPZP-RCS2A-DEST-mRFP-N1 using primers listed in Table 1, clone it by the recombination cloning technique into pDONR207, and then use LR Clonase II to transfer mRFP into pPZP RCS2A-DEST-EGFP-N1 to generate the binary fusion construct p35S::mRFP-GFP (positive control).</li>
	</ol>
	</li>
	<li>Perform transformation of the donor and acceptor constructs into Agrobacterium.
	<ol>
		<li>Add 1 &#181;g of each plasmid from steps 1.2.3-1.2.5 to 100 &#181;L of the culture of competent cells of Agrobacterium tumefaciens strain EHA105 or GV3101, prepared using standard protocols37 or obtained commercially, and incubate at 37 &#176;C for 5 min.</li>
		<li>Add 1 mL of LB medium (1% tryptone, 0.5% yeast extract, and 1% NaCl; see Table of Materials) to the competent cell mixture and agitate at 200 rpm and 28&#176; C for 1.5 h. Collect the cells by centrifugation at 3,000 &#215; g for 1 min at room temperature.</li>
		<li>Resuspend the cells in 0.1 mL of LB medium by pipetting and spread them on LB agar supplemented with the appropriate antibiotics (e.g., 0.01% spectinomycin and 0.005% rifampicin; see Table of Materials). Grow at 28 &#176;C for 2 days.</li>
		<li>Pick individual colonies and inoculate each of them separately into 1 mL of LB broth supplemented with the appropriate antibiotics.</li>
		<li>Grow the cells at 28 &#176;C for 24 h and transfer 0.2 mL of the culture into a new tube. Collect the cells by centrifugation at 10,000 &#215; g for 30 s at room temperature.</li>
		<li>Extract plasmid DNA by a standard protocol for isolating plasmids from Agrobacterium cells38 and resuspend the extracted DNA in 30 &#181;L of water. To identify the colonies harboring the desired plasmids, use 2 &#181;L of the DNA preparation as a template for PCR with gene-specific primers listed in Table 1. Mix 0.7 mL of the identified culture with 0.3 mL of glycerol and store at -80 &#176;C.</li>
	</ol>
	</li>
</ol>
2. Agroinfiltration
<ol>
	<li>Grow Nicotiana benthamiana plants. 
	NOTE: Throughout the entire experiment, all plants must be healthy.
	<ol>
		<li>Sow and grow N. benthamiana seeds in a pot containing wet soil at a high density.</li>
		<li>Keep the planted seeds in a growth chamber set at 23 &#176;C with 16 h of light and 8 h of dark cycle with 150-170 &#181;mol/m2s light intensity until the diameter of the euphyll reaches 0.5 cm.</li>
		<li>Transfer the seedlings to larger pots and allow them to grow in the same chamber with the same parameters. 
		NOTE: Plants are ready for agroinfiltration when their largest leaves are 5-7 cm in diameter, usually within 4-5 weeks. In smaller plants that are too young, the effects of agroinfiltration will be too severe for the FRET analysis.</li>
	</ol>
	</li>
	<li>Prepare bacterial cells for agroinfiltration.
	<ol>
		<li>Grow each Agrobacterium colony containing the FRET constructs overnight in 5 mL of LB medium supplemented with the appropriate antibiotics (step 1.3.3) and 150 &#181;M acetosyringone at 28 &#176;C (see Table of Materials).</li>
		<li>Centrifuge the cells at 3,000 &#215; g for 5 min at room temperature.</li>
		<li>Resuspend the cells in agroinfiltration buffer (10 mM MgCl2, 10 mM MES pH 5.6, 150 &#181;M acetosyringone) to OD600 = 0.5.</li>
		<li>Combine the resuspended cells at a 1:1 v/v ratio between cells harboring the appropriate constructs (step 2.2.5).</li>
		<li>For the double-construct agroinfiltrations, mix the aliquots of two cultures and, for single-construct agroinfiltrations, mix the aliquots of the same culture:
		<ol>
			<li>Tested proteins: OTLD1-mRFP + LSH10-GFP (bacteria harboring the p35S::OTLD1-mRFP and p35S::LSH10-GFP constructs).</li>
			<li>Negative control: OTLD1-mRFP + LSH4-GFP (bacteria harboring the p35S::OTLD1-mRFP and p35S::LSH4-GFP constructs).</li>
			<li>Negative control: LSH10-GFP + free mRFP (bacteria harboring the p35S::LSH10-GFP and pPZP-RCS2A-DEST-mRFP-C1 constructs).</li>
			<li>Positive control: mRFP-GFP (bacteria harboring the p35S::mRFP-GFP construct).</li>
		</ol>
		</li>
		<li>Incubate the cells at 28 &#176;C for 0.5-1 h.</li>
	</ol>
	</li>
	<li>Perform agroinfiltration.
	<ol>
		<li>Load the bacterial culture into a 1 mL needleless syringe.</li>
		<li>Gently but firmly press the nozzle of the syringe against the abaxial side of the fully expanded N. benthamiana leaves while holding the leaf with a gloved finger on the adaxial side.</li>
		<li>Infiltrate up to four spots on a leaf, three leaves per plant, two or three plants for each bacterial culture. Change gloves between samples to prevent cross-contamination.</li>
		<li>Maintain the agroinfiltrated plants in the same growth chamber under the same conditions, as described in step 2.1.2, for 24 h to 36 h. Do not keep the agroinfiltrated plants for longer than 36 h, as this will reduce the fluorescence signal.</li>
	</ol>
	</li>
</ol>
3. Confocal microscopy
<ol>
	<li>Prepare microscope slides for fluorescence visualization.
	<ol>
		<li>After 24-36 h of the infiltration, use a razor blade to cut each agroinfiltrated leaf into small pieces (2 mm x 4 mm) between the veins.</li>
		<li>Place the leaf pieces on a glass slide with the abaxial leaf surface facing up. Place a drop of water on the leaf pieces and cover them with the cover glass. Slightly tap the cover glass to remove air bubbles.</li>
		<li>Turn on the microscope and laser (see Table of Materials). Place the slide into the microscope stage holder for imaging under the specific FRET parameters (steps 3.2 and 3.3).</li>
		<li>Begin the observations using a 10x objective lens to identify cells that exhibit both the GFP and mRFP signals, and then use a 40x objective lens for subsequent detailed observations. 
		NOTE: Importantly, SE-FRET and AB-FRET usually are performed on the same tissue sample, allowing the use of the same channel settings (step 3.2) except for the FRET channel, which is toggled on/off for the SE-FRET and AB-FRET observations, respectively (steps 3.2.2.3 and 3.3.1).</li>
	</ol>
	</li>
	<li>Set up the parameters for SE-FRET (Figure 1A).
	<ol>
		<li>Open the Multi-Dimensional Acquisition (MDA) tool.</li>
		<li>Establish a set of three confocal channels in the same field of view (Supplementary Figure 1).
		<ol>
			<li>Set the donor channel (the GFP channel) for excitation and emission of the donor fluorochrome with the 405 nm excitation laser and 400-597 nm emission filter.</li>
			<li>Set the acceptor channel (the mRFP channel) for excitation and emission of the acceptor fluorochrome with the 561 nm excitation laser and 400-597 nm emission filter. 
			NOTE: The emission filter for mRFP was set at 400-597 nm to separate the mRFP signal from the FRET signal at 597-617 nm (step 3.2.2.3) and, therefore, reduce the FRET-independent mRFP emission.</li>
			<li>Set the FRET channel for excitation of the donor and emission of the acceptor fluorochromes with the 405 nm excitation laser and 597-617 nm emission filter.</li>
		</ol>
		</li>
		<li>Set the donor excitation intensity at a minimum level to observe FRET while avoiding photobleaching, reducing the SE-FRET efficiency. 
		NOTE: This excitation intensity is experimentally selected before conducting the FRET procedure to avoid photobleaching. It varies depending on leaf thickness, age, and time after overexpression.</li>
		<li>Excite the donor and scan for cells containing the acceptor&#39;s expected fluorescence signal.</li>
		<li>Select the region that contains the fluorescence signal of interest.</li>
		<li>Acquire a SE-FRET image sequence by pressing the Snap button.
		<ol>
			<li>Image 10-15 cells expressing the mRFP-GFP construct (positive control) first; adjust the focus, zoom, and smart gain parameters to focus on the area of interest to be captured (Supplementary Figure 2).</li>
			<li>Using the same settings, image 10-15 cells, each expressing OTLD1-mRFP, free mRFP, LSH10-GFP, or LSH4-GFP separately. 
			NOTE: These images are acquired by the &#34;PixFRET&#34; plug-in of ImageJ (see Table of Materials), which was used for the FRET data analyses (step 3.4.1) to determine the spectral bleed-through (SBT) values for the acceptors and the donors; these images are used by the software to generate the SE-FRET images for the OTLD1-mRFP + LSH10-GFP, OTLD1-mRFP + LSH4-GFP, and LSH10-GFP + free mRFP protein pairs (step 3.2.6.3).</li>
			<li>Also, using the same settings, image 10-15 cells co-expressing OTLD1-mRFP + LSH10-GFP, OTLD1-mRFP + LSH4-GFP, and LSH10-GFP + free mRFP protein pairs.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Set up parameters for AB-FRET (Figure 1B).
	<ol>
		<li>Utilize the donor and acceptor channel parameters set for SE-FRET (step 3.2.2) but turn off the FRET channel.</li>
		<li>Set the parameters for photobleaching of the acceptor (mRFP) (Supplementary Figure 3).
		<ol>
			<li>Ensure that bleaching starts after five images. Allow 200 iterations for each area bleach. Keep 100% laser intensity at 561 nm.</li>
			<li>Maintain a bleaching duration of 45 s. Ensure a scan speed of 512 x 512 pixels at 400 Hz.</li>
		</ol>
		</li>
		<li>Draw the region of the cell to be bleached; for example, for nuclear interactions, regions of interest are drawn around the entire area of the cell nucleus39.</li>
		<li>Activate bleaching by pressing the Start experiment button; activating this function will perform the photobleaching and acquire the AB-FRET image sequence.</li>
	</ol>
	</li>
	<li>Analyze the FRET data.
	<ol>
		<li>For analyzing SE-FRET data, use the &#34;PixFRET&#34; plug-in for the ImageJ software to generate corrected images of the SE-FRET efficiency after subtracting SBT40 (step 3.2.6.2).</li>
		<li>For analyzing the AB-FRET data, calculate %AB-FRET as the percent increase in GFP emission after mRFP photobleaching using the following formula41: %AB-FRET = [(GFPpost - GFPpre) / GFPpre] x 100, where GFPpost is GFP emission after mRFP photobleaching, and GFPpre is GFP emission before mRFP photobleaching.</li>
		<li>When reviewing the FRET images, pay attention to the subcellular localization of the FRET signal. 
		NOTE: In many cases, these cellular compartments (e.g., nucleus, chloroplasts, ER, etc.) can be easily identified and, as an additional benefit of the FRET technique, provide important clues to the biological function of the interacting proteins.</li>
	</ol>
	</li>
</ol>

No conflicts of interest were declared.

This FRET protocol is simple and easy to reproduce; it also requires minimal supply investment and utilizes standard equipment for many modern laboratories. Specifically, five main technical features distinguish the versatility of this procedure. First, the FRET constructs are generated using site-specific recombination, a cloning approach that is easy to use, produces accurate results, and saves time compared to traditional restriction enzyme-based cloning. Second, N. benthamiana plants are simple to grow, prod...

Plant histone deubiquitinases play an important role in controlling gene expression by post-translational modification of histones, specifically by erasing their monoubiquitylation marks1. So far, OTLD1 is one of the only few plant histone deubiquitinases characterized at the molecular level in Arabidopsis2,3. OTLD1 removes monoubiquitin groups from the H2B histone molecules, thereby promoting the removal or addition of euchromatic acetylation and methylation modifications of H3 histones in the target gene chromatin4,5. Moreover, OTLD1 interacts with another chromatin-modifying enzyme, the histone lysine demethylase KDM1C, to affect transcriptional suppression of the target genes6,7.
Most histone-modifying enzymes lack DNA binding capabilities, and thus cannot recognize their target genes directly. One possibility is that they cooperate with DNA-binding transcription factor proteins which bind these enzymes and direct them to their chromatin targets. Specifically, in plants, several major histone-modifying enzymes (i.e., histone methyltransferases8,9, histone acetyltransferases10, histone demethylases11, and Polycomb repressive complexes12,13,14) are known to be recruited by transcription factors. Consistent with this idea, recently, one possible mechanism for OTLD1 recruitment to the target promoters was proposed which is based on specific protein-protein interactions of OTLD1 with a transcription factor LSH1015.
LSH10 belongs to a family of the plant ALOG (Arabidopsis LSH1 and Oryza G1) proteins that function as central developmental regulators16,17,18,19,20,21,22. The fact that the members of the ALOG protein family contain DNA binding motifs23 and exhibit the capacities for transcriptional regulation22, nuclear localization19, and homodimerization24 lends further support to the notion that these proteins, including LSH10, may act as specific transcription factors during epigenetic regulation of transcription. One of the main experimental techniques used to characterize the LSH10-OTLD1 interaction in vivo is fluorescence resonance energy transfer (FRET)15.
FRET is an imaging technique for directly detecting close-range interactions between proteins within &#60;10 nm from each other25 inside living cells. There are two main variations of the FRET approach26: sensitized emission (SE-FRET) (Figure&#160;1A) and acceptor bleaching (AB-FRET) (Figure 1B). In SE-FRET, the interacting proteins-one of which is tagged with a donor fluorochrome (e.g., green fluorescent protein, GFP) and the other with an acceptor fluorochrome (e.g., monomeric red fluorescent protein, mRFP27,28)-non-radiatively transfer the excited state energy from the donor to the acceptor. Because no photons are emitted during this transfer, a fluorescent signal is produced that has a radiative emission spectrum similar to that of the acceptor. In AB-FRET, protein interactions are detected and quantified based on elevated radiative emission of the donor when the acceptor is permanently inactivated by photobleaching, and thus is unable to receive the non-radiative energy transferred from the donor (Figure&#160;1). Importantly, the subcellular location of the FRET fluorescence is indicative of the localization of the interacting proteins in the cell.
The ability to deploy FRET in living tissues and determine the subcellular localization of the interacting proteins simultaneously with detecting this interaction per se, makes FRET the technique of choice for studies and initial characterization of protein-protein interactions in vivo. A comparable in vivo fluorescence imaging methodology, bimolecular fluorescence complementation (BiFC)29,30,31,32, is a good alternative approach, although, unlike FRET, BiFC may produce false positives due to spontaneous assembly of the autofluorescent BiFC reporters33, and quantification of its data is less precise.
This article shares the successful experience in implementing both SE-FRET and AB-FRET techniques and presents a protocol for their deployment to investigate the interactions between OTLD1 and LSH10 in plant cells.

<table><tbody><tr><td>Acetosyringone (3&amp;prime;,5&amp;prime;-Dimethoxy-4&amp;prime;-hydroxyacetophenone)</td><td>Sigma-Aldrich</td><td>#D134406-1G</td><td></td></tr><tr><td>Bacto Agar</td><td>BD Biosciences</td><td>#214010</td><td></td></tr><tr><td>Bacto trypton</td><td>BD Biosciences</td><td>#211705</td><td></td></tr><tr><td>Bacto yeast extract</td><td>BD Biosciences</td><td>#212750&amp;nbsp;</td><td></td></tr><tr><td>Confocal laser scanning microscope (CLSM)</td><td>Zeiss</td><td>LSM900</td><td>Any CLSM with similar capabilities is suitable</td></tr><tr><td>EHA105</td><td>VWR</td><td>104013-310</td><td>We use the stock in the Citovsky bacterial lab stock collection</td></tr><tr><td>Gateway BP Clonase II&amp;nbsp;</td><td>Invitrogen</td><td>#11789100</td><td></td></tr><tr><td>Gateway LR Clonase II</td><td>Invitrogen</td><td>#11791020</td><td></td></tr><tr><td>GV3101</td><td>VWR</td><td>104013-296</td><td>We use the stock in the Citovsky bacterial lab stock collection</td></tr><tr><td>ImageJ</td><td></td><td></td><td>https://imagej.nih.gov/ij/download.html</td></tr><tr><td>MES</td><td>Sigma-Aldrich</td><td>#69889-10G</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>#63068-250G</td><td></td></tr><tr><td>NaCl</td><td>Sigma-Aldrich</td><td>#S5886-500G</td><td></td></tr><tr><td>&lt;em&gt;Nicotiana benthamiana&lt;/em&gt; seeds</td><td>Herbalistics Pty</td><td>RA4 or LAB</td><td>We use the stock in the Citovsky seed lab stock collection</td></tr><tr><td>pDONR207</td><td>Invitrogen</td><td>#12213013</td><td></td></tr><tr><td>pPZP-RCS2A-DEST-EGFP-N1&amp;nbsp;</td><td>N/A</td><td></td><td>Refs. 15, 28</td></tr><tr><td>pPZP-RCS2A-DEST-mRFP-C1</td><td>N/A</td><td></td><td>Generated based on the pPZP-RCS2A-DEST-EGFP-C1 construct (see refs. 15, 28)</td></tr><tr><td>pPZP-RCS2A-DEST-mRFP-N1&amp;nbsp;</td><td>N/A</td><td></td><td>Generated based on the pPZP-RCS2A-DEST-EGFP-N1 construct</td></tr><tr><td>Rifampicin</td><td>Sigma-Aldrich</td><td>#R7382-5G</td><td></td></tr><tr><td>Spectinomycin</td><td>Sigma-Aldrich</td><td>#S4014-5G</td><td></td></tr><tr><td>Syringes without needles</td><td>BD</td><td>309659</td><td></td></tr><tr><td>Zen software for CLSM imaging</td><td>Zeiss</td><td>ZEN 3.0 version</td><td>The software should be compatible with the CLSM used</td></tr></tbody></table>

investigating interactions between histone modifying enzymes and transcription factors in vivo by fluorescence resonance energy transfer

Epigenetic regulation of gene expression is commonly affected by histone modifying enzymes (HMEs) that generate heterochromatic or euchromatic histone marks for transcriptional repression or activation, respectively. HMEs are recruited to their target chromatin by transcription factors (TFs). Thus, detecting and characterizing direct interactions between HMEs and TFs are critical for understanding their function and specificity better. These studies would be more biologically relevant if performed in vivo within living tissues. Here, a protocol is described for visualizing interactions in plant leaves between a plant histone deubiquitinase and a plant transcription factor using fluorescence resonance energy transfer (FRET), which allows the detection of complexes between protein molecules that are within &#60;10 nm from each other. Two variations of the FRET technique are presented: SE-FRET (sensitized emission) and AB-FRET (acceptor bleaching), in which the energy is transferred non-radiatively from the donor to the acceptor or emitted radiatively by the donor upon photobleaching of the acceptor. Both SE-FRET and AB-FRET approaches can be adapted easily to discover other interactions between other proteins in planta.

Figure 2 illustrates the typical results of a SE-FRET experiment, in which the cell nuclei were simultaneously recorded in three channels (i.e., donor GFP, acceptor mRFP, and SE-FRET). These data were used to generate images of SE-FRET efficiency coded in a pseudo-color scale. On this scale, the transition from blue to red corresponds to an increase in FRET efficiency, a measure of protein-protein proximity from 0% to 100%. In this representative experiment, the SE-FRET signal was recorded i...

Watch this Scientific Journal Video about Investigating Interactions Between Histone Modifying Enzymes and Transcription Factors in vivo by Fluorescence Resonance Energy Transfer at JoVE.com

Investigating Interactions Between Histone Modifying Enzymes and Transcription Factors in vivo by Fluorescence Resonance Energy Transfer

Fluorescence resonance energy transfer (FRET) is an imaging technique for detecting protein interactions in living cells. Here, a FRET protocol is presented to study the association of histone-modifying enzymes with transcription factors that recruit them to the target promoters for epigenetic regulation of gene expression in plant tissues.

Investigating Interactions Between Histone Modifying Enzymes and Transcription Factors in vivo by Fluorescence Resonance Energy Transfer

Epigenetic regulation of gene expression is commonly affected by histone modifying enzymes (HMEs) that generate heterochromatic or euchromatic histone ...

investigating-interactions-between-histone-modifying-enzymes

Research

JoVE Journal

Biology

1.5K Views.  State University of New York. Fluorescence resonance energy transfer (FRET) is an imaging technique for detecting protein interactions in living cells. Here, a FRET protocol is presented to study the association of histone-modifying enzymes with transcription factors that recruit them to the target promoters for epigenetic regulation of gene expression in plant tissues.

Video: Investigating Interactions Between Histone Modifying Enzymes and Transcription Factors in vivo by Fluorescence Resonance Energy Transfer

Chromatin Immunoprecipitation Assay Using Micrococcal Nucleases in Mammalian Cells

To express cellular phenotypes in organisms, living cells execute gene expression accordingly, and transcriptional programs play a central role in gene expression. The cellular transcriptional machinery and its chromatin modification proteins coordinate to regulate transcription. To analyze transcriptional regulation at the molecular level, several experimental methods such as electrophoretic mobility shift, transient reporter and chromatin immunoprecipitation (ChIP) assays are available. We describe a modified ChIP assay in detail in this article because of its advantages in directly showing histone modifications and the interactions between proteins and DNA in cells. One of the key steps in a successful ChIP assay is chromatin shearing. Although sonication is commonly used for shearing chromatin, it is difficult to identify reproducible conditions. Instead of shearing chromatin by sonication, we utilized enzymatic digestion with micrococcal nuclease (MNase) to obtain more reproducible results. In this article, we provide a straightforward ChIP assay protocol using MNase.

Chromatin immunoprecipitation (ChIP) is a powerful tool for understanding the molecular mechanisms of gene regulation. However, the method involves difficulties in obtaining reproducible chromatin fragmentation by mechanical shearing. Here, we provide an improved protocol for a ChIP assay using enzymatic digestion.

To express cellular phenotypes in organisms, living cells execute gene expression accordingly, and transcriptional programs play a central role in gene ...

Genetics

High Sensitivity Measurement of Transcription Factor-DNA Binding Affinities by Competitive Titration Using Fluorescence Microscopy

Accurate quantification of transcription factor (TF)-DNA interactions is essential for understanding the regulation of gene expression. Since existing approaches suffer from significant limitations, we have developed a new method for determining TF-DNA binding affinities with high sensitivity on a large scale. The assay relies on the established fluorescence anisotropy (FA) principle but introduces important technical improvements. First, we measure a full FA competitive titration curve in a single well by incorporating TF and a fluorescently labeled reference DNA in a porous agarose gel matrix. Unlabeled DNA oligomer is loaded on the top as a competitor and, through diffusion, forms a spatio-temporal gradient. The resulting FA gradient is then read out using a customized epifluorescence microscope setup. This improved setup greatly increases the sensitivity of FA signal detection, allowing both weak and strong binding to be reliably quantified, even for molecules of similar molecular weights. In this fashion, we can measure one titration curve per well of a multi-well plate, and through a fitting procedure, we can extract both the absolute dissociation constant (KD) and active protein concentration. By testing all single-point mutation variants of a given consensus binding sequence, we can survey the entire binding specificity landscape of a TF, typically on a single plate. The resulting position weight matrices (PWMs) outperform those derived from other methods in predicting in vivo TF occupancy. Here, we present a detailed guide for implementing HiP-FA on a conventional automated fluorescent microscope and the data analysis pipeline.

Here we present a novel method for determining binding affinities at equilibrium and in solution with high sensitivity on a large scale. This improves the quantitative analysis of transcription factor-DNA binding. The method is based on automated fluorescence anisotropy measurements in a controlled delivery system.

Accurate quantification of transcription factor (TF)-DNA interactions is essential for understanding the regulation of gene expression. Since existing ...

Biochemistry

Single-Molecule F&ouml;rster Resonance Energy Transfer Methods for Real-Time Investigation of the Holliday Junction Resolution by GEN1

Bulk methods measure the ensemble behavior of molecules, in which individual reaction rates of the underlying steps are averaged throughout the population. Single-molecule F&#246;rster resonance energy transfer (smFRET) provides a recording of the conformational changes taking place by individual molecules in real-time. Therefore, smFRET is powerful in measuring structural changes in the enzyme or substrate during binding and catalysis. This work presents a protocol for single-molecule imaging of the interaction of a four-way Holliday junction (HJ) and gap endonuclease I (GEN1), a cytosolic homologous recombination enzyme. Also presented are single-color and two-color alternating excitation (ALEX) smFRET experimental protocols to follow the resolution of the HJ by GEN1 in real-time. The kinetics of GEN1 dimerization are determined at the HJ, which has been suggested to play a key role in the resolution of the HJ and has remained elusive until now. The techniques described here can be widely applied to obtain valuable mechanistic insights of many enzyme-DNA systems.

Single-Molecule F&#246;rster Resonance Energy Transfer Methods for Real-Time Investigation of the Holliday Junction Resolution by GEN1

Presented here is a protocol for performing single-molecule F&#246;rster resonance energy transfer to study HJ resolution. Two-color alternating excitation is used for determining the dissociation constants. Single-color time lapse smFRET is then applied in real-time cleavage assays to obtain the dwell time distribution prior to HJ resolution.

Bulk methods measure the ensemble behavior of molecules, in which individual reaction rates of the underlying steps are averaged throughout the ...

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 ...

Deciphering Molecular Mechanism of Histone Assembly by DNA Curtain Technique

Chromatin is a higher-order structure that packages eukaryotic DNA. Chromatin undergoes dynamic alterations according to the cell cycle phase and in response to environmental stimuli. These changes are essential for genomic integrity, epigenetic regulation, and DNA metabolic reactions such as replication, transcription, and repair. Chromatin assembly is crucial for chromatin dynamics and is catalyzed by histone chaperones. Despite extensive studies, the mechanisms by which histone chaperones enable chromatin assembly remains elusive. Moreover, the global features of nucleosomes organized by histone chaperones are poorly understood. To address these problems, this work describes a unique single-molecule imaging technique named DNA curtain, which facilitates the investigation of the molecular details of nucleosome assembly by histone chaperones. DNA curtain is a hybrid technique that combines lipid fluidity, microfluidics, and total internal reflection fluorescence microscopy (TIRFM) to provide a universal platform for real-time imaging of diverse protein-DNA interactions.Using DNA curtain, the histone chaperone function of Abo1, the Schizosaccharomyces pombe bromodomain-containing AAA+ ATPase, is investigated, and the molecular mechanism underlying histone assembly of Abo1 is revealed. DNA curtain provides a unique approach for studying chromatin dynamics.

DNA curtain, a high-throughput single-molecule imaging technique, provides a platform for real-time visualization of diverse protein-DNA interactions. The present protocol utilizes the DNA curtain technique to investigate the biological role and molecular mechanism of Abo1, a Schizosaccharomyces pombe bromodomain-containing AAA+ ATPase.

Chromatin is a higher-order structure that packages eukaryotic DNA. Chromatin undergoes dynamic alterations according to the cell cycle phase and in ...

Imaging Protein-protein Interactions in vivo

Protein-protein interactions are a hallmark of all essential cellular processes. However, many of these interactions are transient, or energetically weak, preventing their identification and analysis through traditional biochemical methods such as co-immunoprecipitation. In this regard, the genetically encodable fluorescent proteins (GFP, RFP, etc.) and their associated overlapping fluorescence spectrum have revolutionized our ability to monitor weak interactions in vivo using F&ouml;rster resonance energy transfer (FRET)1-3. Here, we detail our use of a FRET-based proximity assay for monitoring receptor-receptor interactions on the endothelial cell surface.

Imaging Protein-protein Interactions in vivo

This protocol describes how to image protein-protein interactions using a FRET-based proximity assay.

Protein-protein interactions are a hallmark of all essential cellular processes. However, many of these interactions are transient, or energetically weak, ...

Repressing Gene Transcription by Redirecting Cellular Machinery with Chemical Epigenetic Modifiers

Regulation of chromatin compaction is an important process that governs gene expression in higher eukaryotes. Although chromatin compaction and gene expression regulation are commonly disrupted in many diseases, a locus-specific, endogenous, and reversible method to study and control these mechanisms of action has been lacking. To address this issue, we have developed and characterized novel gene-regulating bifunctional molecules. One component of the bifunctional molecule binds to a DNA-protein anchor so that it will be recruited to an allele-specific locus. The other component engages endogenous cellular chromatin-modifying machinery, recruiting these proteins to a gene of interest. These small molecules, called chemical epigenetic modifiers (CEMs), are capable of controlling gene expression and the chromatin environment in a dose-dependent and reversible manner. Here, we detail a CEM approach and its application to decrease gene expression and histone tail acetylation at a Green Fluorescent Protein (GFP) reporter located at the Oct4 locus in mouse embryonic stem cells (mESCs). We characterize the lead CEM (CEM23) using fluorescent microscopy, flow cytometry, and chromatin immunoprecipitation (ChIP), followed by a quantitative polymerase chain reaction (qPCR). While the power of this system is demonstrated at the Oct4 locus, conceptually, the CEM technology is modular and can be applied in other cell types and at other genomic loci.

Regulation of the chromatin environment is an essential process required for proper gene expression. Here, we describe a method for controlling gene expression through the recruitment of chromatin-modifying machinery in a gene-specific and reversible manner.

Regulation of chromatin compaction is an important process that governs gene expression in higher eukaryotes. Although chromatin compaction and gene ...

Bioengineering

Investigating Protein-protein Interactions in Live Cells Using Bioluminescence Resonance Energy Transfer

Assays based on Bioluminescence Resonance Energy Transfer (BRET) provide a sensitive and reliable means to monitor protein-protein interactions in live cells. BRET is the non-radiative transfer of energy from a 'donor' luciferase enzyme to an 'acceptor' fluorescent protein. In the most common configuration of this assay, the donor is Renilla reniformis luciferase and the acceptor is Yellow Fluorescent Protein (YFP). Because the efficiency of energy transfer is strongly distance-dependent, observation of the BRET phenomenon requires that the donor and acceptor be in close proximity. To test for an interaction between two proteins of interest in cultured mammalian cells, one protein is expressed as a fusion with luciferase and the second as a fusion with YFP. An interaction between the two proteins of interest may bring the donor and acceptor sufficiently close for energy transfer to occur. Compared to other techniques for investigating protein-protein interactions, the BRET assay is sensitive, requires little hands-on time and few reagents, and is able to detect interactions which are weak, transient, or dependent on the biochemical environment found within a live cell. It is therefore an ideal approach for confirming putative interactions suggested by yeast two-hybrid or mass spectrometry proteomics studies, and in addition it is well-suited for mapping interacting regions, assessing the effect of post-translational modifications on protein-protein interactions, and evaluating the impact of mutations identified in patient DNA.

Interactions between proteins are fundamental to all cellular processes. Using Bioluminescence Resonance Energy Transfer, the interaction between a pair of proteins can be monitored in live cells and in real time. Furthermore, the effects of potentially pathogenic mutations can be assessed.

Assays based on Bioluminescence Resonance Energy Transfer (BRET) provide a sensitive and reliable means to monitor protein-protein interactions in live ...

Assessing Protein Interactions in Live-Cells with FRET-Sensitized Emission

F&#246;rster Resonance Energy Transfer (FRET) is the radiationless transfer of energy from an excited donor to an acceptor molecule and depends upon the distance and orientation of the molecules as well as the extent of overlap between the donor emission and acceptor absorption spectra. FRET permits to study the interaction of proteins in the living cell over time and in different subcellular compartments. Different intensity-based algorithms to measure FRET using microscopy have been described in the literature. Here, a protocol and an algorithm are provided to quantify FRET efficiency based on measuring both the sensitized emission of the acceptor and quenching of the donor molecule. The quantification of ratiometric FRET in the living cell not only requires the determination of the crosstalk (spectral spill-over, or bleed-through) of the fluorescent proteins but also the detection efficiency of the microscopic setup. The protocol provided here details how to assess these critical parameters.

F&#246;rster Resonance Energy Transfer (FRET) between two fluorophore molecules can be used for studying protein interactions in the living cell. Here, a protocol is provided as to how to measure FRET in live cells by detecting sensitized emission of the acceptor and quenching of the donor molecule using confocal laser scanning microscopy.

F&#246;rster Resonance Energy Transfer (FRET) is the radiationless transfer of energy from an excited donor to an acceptor molecule and depends upon the ...

Determination of Tripartite Interaction between Two Monomers of a MADS-box Transcription Factor and a Calcium Sensor Protein by BiFC-FRET-FLIM Assay

Protein-protein interactions are an integral part of all biological processes in the cells as they play a crucial role in regulating, maintaining, and amending cellular functions. These interactions are involved in a wide range of phenomena such as signal transduction, pathogen response, cell-cell interactions, metabolic and developmental processes. In the case of transcription factors, these interactions may lead to oligomerization of subunits, sequestering in specific subcellular contexts such as the nucleus, cytoplasm, etc., which, in turn, might have a more profound effect on the expression of the downstream genes. Here, we demonstrate a methodology to visualize in vivo tripartite interaction using Bimolecular Fluorescence Complementation (BiFC) based F&#246;rster Resonance Energy Transfer (FRET) involving Fluorescence Lifetime Imaging (FLIM). Two of the proteins selected for this demonstration interact as BiFC partners, and their reconstituted fluorescence activity is used to assay FRET-FLIM with the third partner. Four to five-week-old growth-chamber-grown Nicotiana benthamiana plants have been used as the model plant system for this demonstration.

Here we present, a method to visualize ternary complex formation between three protein partners using fluorescent-tagged proteins by BiFC based FRET-FLIM assay. This method is valuable for studying protein-protein interaction complexes in vivo.

Protein-protein interactions are an integral part of all biological processes in the cells as they play a crucial role in regulating, maintaining, and ...

Investigating Interactions Between Histone Modifying Enzymes and Transcription Factors in vivo by Fluorescence Resonance Energy Transfer

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Chromatin Immunoprecipitation Assay Using Micrococcal Nucleases in Mammalian Cells

High Sensitivity Measurement of Transcription Factor-DNA Binding Affinities by Competitive Titration Using Fluorescence Microscopy

Single-Molecule Förster Resonance Energy Transfer Methods for Real-Time Investigation of the Holliday Junction Resolution by GEN1

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

Deciphering Molecular Mechanism of Histone Assembly by DNA Curtain Technique

Imaging Protein-protein Interactions in vivo

Repressing Gene Transcription by Redirecting Cellular Machinery with Chemical Epigenetic Modifiers

Investigating Protein-protein Interactions in Live Cells Using Bioluminescence Resonance Energy Transfer

Assessing Protein Interactions in Live-Cells with FRET-Sensitized Emission

Determination of Tripartite Interaction between Two Monomers of a MADS-box Transcription Factor and a Calcium Sensor Protein by BiFC-FRET-FLIM Assay