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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Acetosyringone (3&#8242;,5&#8242;-Dimethoxy-4&#8242;-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&#160;</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&#160;</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>MgCl2</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>Nicotiana benthamiana 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&#160;</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&#160;</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 ...

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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

Investigating Tissue- and Organ-specific Phytochrome Responses using FACS-assisted Cell-type Specific Expression Profiling in Arabidopsis thaliana

Light mediates an array of developmental and adaptive processes throughout the life cycle of a plant. Plants utilize light-absorbing molecules called photoreceptors to sense and adapt to light. The red/far-red light-absorbing phytochrome photoreceptors have been studied extensively. Phytochromes exist as a family of proteins with distinct and overlapping functions in all higher plant systems in which they have been studied1. Phytochrome-mediated light responses, which range from seed germination through flowering and senescence, are often localized to specific plant tissues or organs2. Despite the discovery and elucidation of individual and redundant phytochrome functions through mutational analyses, conclusive reports on distinct sites of photoperception and the molecular mechanisms of localized pools of phytochromes that mediate spatial-specific phytochrome responses are limited. We designed experiments based on the hypotheses that specific sites of phytochrome photoperception regulate tissue- and organ-specific aspects of photomorphogenesis, and that localized phytochrome pools engage distinct subsets of downstream target genes in cell-to-cell signaling. We developed a biochemical approach to selectively reduce functional phytochromes in an organ- or tissue-specific manner within transgenic plants. Our studies are based on a bipartite enhancer-trap approach that results in transactivation of the expression of a gene under control of the Upstream Activation Sequence (UAS) element by the transcriptional activator GAL43. The biliverdin reductase (BVR) gene under the control of the UAS is silently maintained in the absence of GAL4 transactivation in the UAS-BVR parent4. Genetic crosses between a UAS-BVR transgenic line and a GAL4-GFP enhancer trap line result in specific expression of the BVR gene in cells marked by GFP expression4. BVR accumulation in Arabidopsis plants results in phytochrome chromophore deficiency in planta5-7. Thus, transgenic plants that we have produced exhibit GAL4-dependent activation of the BVR gene, resulting in the biochemical inactivation of phytochrome, as well as GAL4-dependent GFP expression. Photobiological and molecular genetic analyses of BVR transgenic lines are yielding insight into tissue- and organ-specific phytochrome-mediated responses that have been associated with corresponding sites of photoperception4, 7, 8. Fluorescence Activated Cell Sorting (FACS) of GFP-positive, enhancer-trap-induced BVR-expressing plant protoplasts coupled with cell-type-specific gene expression profiling through microarray analysis is being used to identify putative downstream target genes involved in mediating spatial-specific phytochrome responses. This research is expanding our understanding of sites of light perception, the mechanisms through which various tissues or organs cooperate in light-regulated plant growth and development, and advancing the molecular dissection of complex phytochrome-mediated cell-to-cell signaling cascades.

Investigating Tissue- and Organ-specific Phytochrome Responses using FACS-assisted Cell-type Specific Expression Profiling in Arabidopsis thaliana

The molecular basis of spatial-specific phytochrome responses is being investigated using transgenic plants that exhibit tissue- and organ-specific phytochrome deficiencies. The isolation of specific cells exhibiting induced phytochrome chromophore depletion by Fluorescence-Activated Cell Sorting followed by microarray analyses is being utilized to identify genes involved in spatial-specific phytochrome responses.

Light mediates an array of developmental and adaptive processes throughout the life cycle of a plant. Plants utilize light-absorbing molecules called ...

Gramicidin-based Fluorescence Assay; for Determining Small Molecules Potential for Modifying Lipid Bilayer Properties

Many drugs and other small molecules used to modulate biological function are amphiphiles that adsorb at the bilayer/solution interface and thereby alter lipid bilayer properties. This is important because membrane proteins are energetically coupled to their host bilayer by hydrophobic interactions. Changes in bilayer properties thus alter membrane protein function, which provides an indirect way for amphiphiles to modulate protein function and a possible mechanism for "off-target" drug effects. We have previously developed an electrophysiological assay for detecting changes in lipid bilayer properties using linear gramicidin channels as probes 3,12. Gramicidin channels are mini-proteins formed by the transbilayer dimerization of two non-conducting subunits. They are sensitive to changes in their membrane environment, which makes them powerful probes for monitoring changes in lipid bilayer properties as sensed by bilayer spanning proteins. We now demonstrate a fluorescence assay for detecting changes in bilayer properties using the same channels as probes. The assay is based on measuring the time-course of fluorescence quenching from fluorophore-loaded large unilamellar vesicles due to the entry of a quencher through the gramicidin channels. We use the fluorescence indicator/quencher pair 8-aminonaphthalene-1,3,6-trisulfonate (ANTS)/Tl+ that has been successfully used in other fluorescence quenching assays 5,13. Tl+ permeates the lipid bilayer slowly 8 but passes readily through conducting gramicidin channels 1,14. The method is scalable and suitable for both mechanistic studies and high-throughput screening of small molecules for bilayer-perturbing, and potential "off-target", effects. We find that results using this method are in good agreement with previous electrophysiological results 12.

We introduce a fast fluorescence-based assay that monitors the rate of fluorescence quenching as a measure of gramicidin channel activity. The gramicidin channels are used as molecular force transducers to monitor changes in lipid bilayer properties as sensed by bilayer spanning proteins.

Many drugs and other small molecules used to modulate biological function are amphiphiles that adsorb at the bilayer/solution interface and thereby alter ...

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, ...

Real-time Monitoring of Ligand-receptor Interactions with Fluorescence Resonance Energy Transfer

FRET is a process whereby energy is non-radiatively transferred from an excited donor molecule to a ground-state acceptor molecule through long-range dipole-dipole interactions1. In the present sensing assay, we utilize an interesting property of PDA: blue-shift in the UV-Vis electronic absorption spectrum of PDA (Figure 1) after an analyte interacts with receptors attached to PDA2,3,4,7. This shift in the PDA absorption spectrum provides changes in the spectral overlap (J) between PDA (acceptor) and rhodamine (donor) that leads to changes in the FRET efficiency. Thus, the interactions between analyte (ligand) and receptors are detected through FRET between donor fluorophores and PDA. In particular, we show the sensing of a model protein molecule streptavidin. We also demonstrate the covalent-binding of bovine serum albumin (BSA) to the liposome surface with FRET mechanism. These interactions between the bilayer liposomes and protein molecules can be sensed in real-time. The proposed method is a general method for sensing small chemical and large biochemical molecules. Since fluorescence is intrinsically more sensitive than colorimetry, the detection limit of the assay can be in sub-nanomolar range or lower8. Further, PDA can act as a universal acceptor in FRET, which means that multiple sensors can be developed with PDA (acceptor) functionalized with donors and different receptors attached on the surface of PDA liposomes.

We demonstrate FRET between conjugated polymer polydiacetylene (PDA) and fluorophore attached to the surface of PDA liposomes for the sensing of biomolecules. PDA liposomes also contained receptor molecules on their surfaces for biomolecules to be used as probes. Ligand-receptor interactions lead to changes in the FRET efficiency between the fluorophore and PDA which is the basis of the sensing mechanism.

FRET is a process  whereby energy is non-radiatively transferred from an excited donor molecule to  a ground-state acceptor molecule through long-range ...

In vivo Reprogramming of Adult Somatic Cells to Pluripotency by Overexpression of Yamanaka Factors

Induced pluripotent stem (iPS) cells that result from the reprogramming of somatic cells to a pluripotent state by forced expression of defined factors are offering new opportunities for regenerative medicine. Such clinical applications of iPS cells have been limited so far, mainly due to the poor efficiency of the existing reprogramming methodologies and the risk of the generated iPS cells to form tumors upon implantation.
We hypothesized that the reprogramming of somatic cells towards pluripotency could be achieved in vivo by gene transfer of reprogramming factors. In order to efficiently reprogram cells in vivo, high levels of the Yamanaka (OKSM) transcription factors need to be expressed at the target tissue. This can be achieved by using different viral or nonviral gene vectors depending on the target tissue. In this particular study, hydrodynamic tail-vein (HTV) injection of plasmid DNA was used to deliver the OKSM factors to mouse hepatocytes. This provided proof-of-evidence of in vivo reprogramming of adult, somatic cells towards a pluripotent state with high efficiency and fast kinetics. Furthermore no tumor or teratoma formation was observed in situ.
It can be concluded that reprogramming somatic cells in vivo may offer a potential approach to induce enhanced pluripotency rapidly, efficiently, and safely compared to in vitro performed protocols and can be applied to different tissue types in the future.

In vivo Reprogramming of Adult Somatic Cells to Pluripotency by Overexpression of Yamanaka Factors

This study demonstrates the reprogramming of somatic cells towards pluripotency in vivo without the generation of teratomas. We used hydrodynamic tail vein injection of plasmid DNA encoding the Yamanka factors to induce the in vivo reprogramming of adult hepatocytes into cells of enhanced pluripotency.

Induced pluripotent stem (iPS) cells that result from the reprogramming of somatic cells to a pluripotent state by forced expression of defined factors ...

Rapid Synthesis and Screening of Chemically Activated Transcription Factors with GFP-based Reporters

Synthetic biology aims to rationally design and build synthetic circuits with desired quantitative properties, as well as provide tools to interrogate the structure of native control circuits. In both cases, the ability to program gene expression in a rapid and tunable fashion, with no off-target effects, can be useful. We have constructed yeast strains containing the ACT1 promoter upstream of a URA3 cassette followed by the ligand-binding domain of the human estrogen receptor and VP16. By transforming this strain with a linear PCR product containing a DNA binding domain and selecting against the presence of URA3, a constitutively expressed artificial transcription factor (ATF) can be generated by homologous recombination. ATFs engineered in this fashion can activate a unique target gene in the presence of inducer, thereby eliminating both the off-target activation and nonphysiological growth conditions found with commonly used conditional gene expression systems. A simple method for the rapid construction of GFP reporter plasmids that respond specifically to a native or artificial transcription factor of interest is also provided.

This protocol describes an experimental procedure for the rapid construction of artificial transcription factors (ATFs) with cognate GFP reporters&#160;and quantification of the ATFs ability to stimulate GFP expression via flow cytometry.

Synthetic biology aims to rationally design and build synthetic circuits with desired quantitative properties, as well as provide tools to interrogate the ...

Protein-protein Interactions Visualized by Bimolecular Fluorescence Complementation in Tobacco Protoplasts and Leaves

Many proteins interact transiently with other proteins or are integrated into multi-protein complexes to perform their biological function. Bimolecular fluorescence complementation (BiFC) is an in vivo method to monitor such interactions in plant cells. In the presented protocol the investigated candidate proteins are fused to complementary halves of fluorescent proteins and the respective constructs are introduced into plant cells via agrobacterium-mediated transformation. Subsequently, the proteins are transiently expressed in tobacco leaves and the restored fluorescent signals can be detected with a confocal laser scanning microscope in the intact cells. This allows not only visualization of the interaction itself, but also the subcellular localization of the protein complexes can be determined. For this purpose, marker genes containing a fluorescent tag can be coexpressed along with the BiFC constructs, thus visualizing cellular structures such as the endoplasmic reticulum, mitochondria, the Golgi apparatus or the plasma membrane. The fluorescent signal can be monitored either directly in epidermal leaf cells or in single protoplasts, which can be easily isolated from the transformed tobacco leaves. BiFC is ideally suited to study protein-protein interactions in their natural surroundings within the living cell. However, it has to be considered that the expression has to be driven by strong promoters and that the interaction partners are modified due to fusion of the relatively large fluorescence tags, which might interfere with the interaction mechanism. Nevertheless, BiFC is an excellent complementary approach to other commonly applied methods investigating protein-protein interactions, such as coimmunoprecipitation, in vitro pull-down assays or yeast-two-hybrid experiments.

Formation of protein complexes in vivo can be visualized by bimolecular fluorescence complementation. Interaction partners are fused to complementary parts of fluorescent tags and transiently expressed in tobacco leaves, resulting in a reconstituted fluorescent signal upon close proximity of the two proteins.

Many proteins interact transiently with other proteins or are integrated into multi-protein complexes to perform their biological function. Bimolecular ...

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 ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

Functional Imaging of Viral Transcription Factories Using 3D Fluorescence Microscopy

It is well known that spatial and temporal regulation of genes is an integral part of governing proper gene expression. Consequently, it is invaluable to understand where and when transcription is taking place within nuclear space and to visualize the relationship between episomes infected within the same cell's nucleus. Here, both immunofluorescence (IFA) and RNA-FISH have been combinedto identify actively transcribing Kaposi's sarcoma-associated herpesvirus (KSHV) episomes. By staining KSHV latency-associated nuclear antigen (LANA), it is possible to locate where viral episomes exist within the nucleus. In addition, by designing RNA-FISH probes to target the intron region of a viral gene, which is expressed only during productive infection, nascent RNA transcripts can be located. Using this combination of molecular probes, it is possible to visualize the assembly of large viral transcription factories and analyze the spatial regulation of viral gene expression during KSHV reactivation. By including anti-RNA polymerase II antibody staining, one can also visualize the association between RNA polymerase II (RNAPII) aggregation and KSHV transcription during reactivation.

Viral transcriptional factories are discrete structures that are enriched with cellular RNA polymerase II to increase viral gene transcription during reactivation. Here, a method to locate sites of actively transcribing viral chromatin in 3D nuclear space by a combination of immunofluorescence staining and in situ RNA hybridization is described.

It is well known that spatial and temporal regulation of genes is an integral part of governing proper gene expression. Consequently, it is invaluable to ...