Name: imaging protein-protein interactions in vivo
Uploaded: 2010-10-10T17:00:00
Duration: 11 min 15 s
Description: Watch this Scientific Journal Video about Imaging Protein-protein Interactions in vivo at JoVE.com

We wish to acknowledge Dr Scott Henderson for help with confocal microscopy. This research was supported by grants from the National Institutes of Health 1RO1CA127501 to W.A.B as well as pilot project funding from the Massey Cancer Center and
School of Medicine (VCU) to W.A.B. Microscopy was performed at the VCU-Dept. of Neurobiology &amp; Anatomy Microscopy Facility, supported, in part, with funding from NIHNINDS Center core grant 5P30NS047463.

The protocol consists of three major steps. The first, step cloning of your gene of interest into a mammalian expression vector amino-terminally to the monomeric versions of CFP and/or YFP will only be discussed briefly. The second and third steps; transfection of vector DNA into EA.hy926 endothelial cells, and confocal imaging with FRET, are outlined in more depth below.
1. Construction of CFP and YFP Chimeric Receptors
Receptors of interest should be cloned amino-terminally to monomeric enhanced versions of CFP and YFP. Determination of FRET is empirical and is critically dependent upon the length of linker between the transmembrane spanning region and start of the fluorophore1. Therefore, several different lengths of linker must be explored for each new receptor pair under investigation.
<ol>
<li> Into the pcDNA3.1 hygroR or neoR vector containing monomeric C- or YFP (this vector can be obtained from our lab), clone your receptor of interest into the NheI/EcoRI sites. *Note-If necessary, NheI is compatible several other enzymes including SpeI and XbaI.</li>
<li> Transform your ligation reaction into an appropriate endA1 recA competent cell line (we routinely use Mach1). Screen colonies for the presence of insert followed by DNA sequencing to verify the absence of mutations.</li>
<li> Upon obtaining conformation of successful molecular cloning, prepare transfection-grade vector DNA in sterilized water or TE using Qiagen, or similar, DNA kits. Evaluate concentration and purity using UV spectroscopy. Typical yields of DNA should be in the range of 100-200 &mu;g, which is routinely diluted to a working stock of 1 &mu;g/&mu;L.</li>
</ol>
2. Transfection of EA.hy 926 Cells
<ol>
<li> Split EA.hy 926 cells4, grown in complete-DMEM (DMEM+10% Fetal bovine serum+Pen./Strep.) into MatTek coverslip (No. 0) 35 mm dishes with 2 mL medium. You can conservatively obtain ~15 dishes (~90% confluent the following day) from a 15 cm confluent dish. Prepare one dish to evaluate expression of each individual receptor, as well as one dish for each receptor pair.</li>
<li> Allow the cells to incubate overnight in 5%CO2 atmosphere at 37&deg;C. The following day, the cultures should be ~ 70-90% confluent.</li>
<li> On the day of transfection, prepare the nucleolipid complexes by mixing 2 &mu;g of the chimeric receptor vector DNA (2 &mu;L) with 8 &mu;L of Fugene6 in 200 &mu;L of pre-warmed OptiMEM. For receptor pairs, transfect with 1 &mu;g of each receptor, for a total of 2 &mu;g (See also discussion below).</li>
<li> Mix gently by inversion.</li>
<li> Continue to incubate at room temperature for 30 minutes.</li>
<li> During the incubation period, prepare the cells for transfection. Aspirate off fresh media and wash each plate gently and briefly with warm PBS before the addition of 2 mL pre-warmed complete-DMEM without antibiotics (DMEM+FBS). It is important to avoid the presence of Penicillin and Streptomycin, as they are considerably more toxic to cells during treatment with Fugene6.</li>
<li> Add the complex drop wise to cells and incubate for an additional 20-24 hours at 37&deg;C in a 5% CO2 atmosphere.</li>
<li> After 24 hours, carefully aspirate the media from the cells and pipette fresh complete-DMEM with antibiotics. Following transfection, cell viability should remain relatively high, i.e. little to no floating cells should be observed.</li>
<li> Gene expression is typically maximal after an additional 48-60 hours post-transfection. However, imaging is typically feasible in as short as 24 hours. Time course experiments for optimal gene expression should be performed.</li>
</ol>
3. Confocal Imaging and FRET Analysis
Live cell imaging is performed on a Leica TCS-SP2 AOBS confocal laser scanning microscope equipped with blue diode (405 nm), Argon (458, 476, 488, 514 nm). green HeNe (543 nm), orange HeNe (594 nm), and red HeNe (633 nm) lasers, an HCX PI Apo 63x/1.3 n.a. glycerin-immersion objective lens, a motorized XY stage (M&auml;rzh&auml;user), and an environmentally controlled (temperature, humidity, and CO2) stage incubator (PeCon). Experimental controls are critical to eliminate fluorophore cross-talk between the emission channels as well as to evaluate expression issues and non-specific fluorophore multimerization. As such, single fluorophore transfections are utilized for setting up every image session. Negative FRET controls (using known noninteracting receptors) will need to be performed, in order to determine the background FRET that occurs due to over expression.
<ol>
<li> To eliminate crosstalk between the emission channels for CFP and YFP, place the CFP expression control into the stage incubator. Using white light, adjust the Z-plate to focus on to the cells.</li>
<li> Set SP window settings for emission detection of CFP to 465-505 nm and YFP to 525-600 nm.</li>
<li> While monitoring both emission channels, excite the CFP at 458 nm. Using the AOTF, set the laser power to eliminate appreciable bleed over of CFP into the YFP emission channel. Save this setting.</li>
<li> Perform the same control for YFP crosstalk (using the YFP expression control) in the CFP channel by adjusting the excitation power at 514 nm for cells expressing only YFP. Save this setting.</li>
<li> Next place cells co-expressing CFP and YFP into the stage incubator. Using the sequence setting for the Leica confocal software, locate cells expressing similar levels of CFP and YFP with proper membrane localization.</li>
<li> Using the Leica software acceptor photo-bleaching program, set the donor and acceptor settings to your saved CFP and YFP settings, respectably.</li>
<li> Zooming onto the cell, highlight an ROI (region of interest) in which the photo-destruction of YFP is to occur on the cellular membrane and begin the program.</li>
<li> For photo-destruction of the acceptor, adjust the AOTF to 100% at 514 nm and allow the laser bleaching to continue until 70% of the YFP emission has been depleted. To accelerate bleaching, ROIs are typically chosen in the raster axis of the laser (X-axis).</li>
<li> During photo-bleaching, monitor the cellular movement and reject measurements obtained with appreciable movement in the XY-plane, as these measurements can report false FRET efficiencies.</li>
<li> Pre- and post-bleach images are recorded for both donor (CFP) and acceptor (YFP) and FRET efficiency is calculated as: FRETEff=(Dpost-Dpre)/Dpost for all Dpost &gt; Dpre where Dpre and Dpost is the donor intensity before and after photobleaching respectively.</li>
<li> JMP Software (SAS) is used to determine the extent of experimental variability.</li>
</ol>
4. Representative Results
Transfection efficiencies are usually between 30 to 40%. Despite previous findings by others, we have not seen that transfection and expression of one vector favors that of the other. Indeed, we frequently observe exclusive expression of one fluorophore chimera or the other. Typical FRET efficiencies vary among receptor systems. For the Tie receptor system, typical values are 20-28% for epithelial cells and 19-23% in endothelial cells. For negative controls, typical efficiencies are below 2-3%. The extent of variability will decrease considerably with experience.
<img src="/files/ftp_upload/2149/2149fig1.jpg" alt="Figure 1" /> 
Figure 1. Representative images of transfected EA.hy 926 cells. A) DIC image of EA.hy 926 cell monolayer. B) Fluorescence image of cells displayed in (A). C) Overlay of (A) and (B) demonstrating ~20-30% transfection efficiency.
<img src="/files/ftp_upload/2149/2149fig2.jpg" alt="Figure 2" /> 
Figure 2. Representative acceptor photo-bleaching analysis of FRET occurring between CFP and YFP in an EA.hy926 cell. The CFP emission channel (top panels) and YFP emission channel (bottom two panels) were monitored separately prior to and post acceptor photobleaching. Photobleaching experiments were restricted to, and FRET values calculated from, the region within the green box. FRET efficiency is displayed as an absolute range from high (red-1.0) to low (purple-0.0) on a magnified overlay of a CFP/YFP merged image for orientation purposes only.

<ol>
	<li>Kim, M., Carman, C. V., Springer, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bidirectional+transmembrane+signaling+by+cytoplasmic+domain+separation+in+integrins.">Bidirectional transmembrane signaling by cytoplasmic domain separation in integrins.</a> Science. 301, 1720-1725 (2003).</li><li>Zacharias, D. A., Violin, J. D., Newton, A. C., Tsien, R. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Partitioning+of+lipidmodified+monomeric+GFPs+into+membrane+microdomains+of+live+cells.">Partitioning of lipidmodified monomeric GFPs into membrane microdomains of live cells.</a> Science. 296, 913-916 (2002).</li><li>Seegar, T. C. M., Eller, B., Tzvetkova-Robev, D., Kolev, M., Henderson, S. C., Nikolov, D. B., Barton, W. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tie1-Tie2+interactions+mediate+functional+differences+between+angiopoietin+ligands.">Tie1-Tie2 interactions mediate functional differences between angiopoietin ligands.</a> Molecular Cell. 37 (5), 643-655 (2010).</li><li>Edgell, C. J., McDonald, C. C., Graham, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Permanent+cell+line+expressing+human+factor+VIII-related+antigen+established+by+hybridization.">Permanent cell line expressing human factor VIII-related antigen established by hybridization.</a> Proc Natl Acad Sci U S A. 80 (12), 3734-3737 (1983).</li></ol>

No conflicts of interest declared.

There are several steps that are critical to success. The most prominent among them is the relative levels of expression between the two chimeric receptors. To circumvent this issue, one may make stable cell lines expressing both proteins of interest, or identify optimal ratios of vector DNA to permit equivalent expression. Similarly, due to non-uniform transfection across a dish, protein levels will seldom be equivalent among cells. Therefore, attention must be paid to distinguish 'high' expressors from 'low'. Those that display 'average' levels of protein typically yield reliable and reproducible FRET efficiencies. One method our lab has pursued to relieve this issue is the use of adeno and lenti-viruses to 'even' gene expression. Furthermore, an alternative method to acceptor photo-bleaching for determining FRET efficiencies is sensitized emission. Although, despite the potential of sensitized emission to monitor single cells in real-time, we have found acceptor photo-bleaching to be more sensitive and more reliable.
Finally, using FRET to monitor protein interactions can be challenging, and requires careful selection of linker lengths for membrane bound receptors. Furthermore, addition of C/YFP often greatly influences protein expression levels and results in aggregation in the endoplasmic reticulum and golgi apparatus. However, for transient, or unstable, interactions, FRET is the ideal methodology to utilize.

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<td>DMEM</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>11960-069</td>
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<td>Penicillin- Streptomycin</td>
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<td>Invitrogen</td>
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<td>Hyclone</td>
<td>N/A</td>
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<td>Opti-MEM</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
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<td>FUGENE 6</td>
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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.

Watch this Scientific Journal Video about Imaging Protein-protein Interactions in vivo at JoVE.com

Imaging Protein-protein Interactions in vivo

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

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, ...

imaging-protein-protein-interactions-in-vivo

Research

JoVE Journal

Biology

Split-Ubiquitin Based Membrane Yeast Two-Hybrid (MYTH) System: A Powerful Tool For Identifying Protein-Protein Interactions

The fundamental biological and clinical importance of integral membrane proteins prompted the development of a yeast-based system for the high-throughput identification of protein-protein interactions (PPI) for full-length transmembrane proteins. To this end, our lab developed the split-ubiquitin based Membrane Yeast Two-Hybrid (MYTH) system. This technology allows for the sensitive detection of transient and stable protein interactions using Saccharomyces cerevisiae as a host organism. MYTH takes advantage of the observation that ubiquitin can be separated into two stable moieties: the C-terminal half of yeast ubiquitin (Cub) and the N-terminal half of the ubiquitin moiety (Nub). In MYTH, this principle is adapted for use as a 'sensor' of protein-protein interactions. Briefly, the integral membrane bait protein is fused to Cub which is linked to an artificial transcription factor. Prey proteins, either in individual or library format, are fused to the Nub moiety. Protein interaction between the bait and prey leads to reconstitution of the ubiquitin moieties, forming a full-length 'pseudo-ubiquitin' molecule. This molecule is in turn recognized by cytosolic deubiquitinating enzymes, resulting in cleavage of the transcription factor, and subsequent induction of reporter gene expression. The system is highly adaptable, and is particularly well-suited to high-throughput screening. It has been successfully employed to investigate interactions using integral membrane proteins from both yeast and other organisms.

Split-Ubiquitin Based Membrane Yeast Two-Hybrid MYTH System: A Powerful Tool For Identifying Protein-Protein Interactions

MYTH allows the sensitive detection of transient and stable interactions between proteins that are expressed in the model organism Saccharomyces cerevisiae. It has been successfully applied to study exogenous and yeast integral membrane proteins in order to identify their interacting partners in a high throughput manner.

The fundamental biological and clinical importance of integral membrane proteins prompted the development of a yeast-based system for the high-throughput ...

In-vivo Detection of Protein-protein Interactions on Micro-patterned Surfaces

Unraveling the interaction network of molecules in-vivo is key to understanding the mechanisms that regulate cell function and metabolism. A multitude of methodological options for addressing molecular interactions in cells have been developed, but most of these methods suffer from being rather indirect and therefore hardly quantitative. On the contrary, a few high-end quantitative approaches were introduced, which however are difficult to extend to high throughput. To combine high throughput capabilities with the possibility to extract quantitative information, we recently developed a new concept for identifying protein-protein interactions (Schwarzenbacher et al., 2008). Here, we describe a detailed protocol for the design and the construction of this system which allows for analyzing interactions between a fluorophore-labeled protein ("prey") and a membrane protein ("bait") in-vivo. Cells are plated on micropatterned surfaces functionalized with antibodies against the bait exoplasmic domain. Bait-prey interactions are assayed via the redistribution of the fluorescent prey. The method is characterized by high sensitivity down to the level of single molecules, the capability to detect weak interactions, and high throughput capability, making it applicable as screening tool.

In-vivo Detection of Protein-protein Interactions on Micro-patterned Surfaces

This video shows experiments with subsequent analysis of protein-protein interactions by the use of micro-patterned surfaces. The approach offers the possibility to detect protein interactions in living cells and combines high throughput capabilities with the possibility to extract quantitative information.

Unraveling the interaction network of molecules in-vivo is key to understanding the mechanisms that regulate cell function and metabolism. A multitude of ...

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) ...

Probing High-density Functional Protein Microarrays to Detect Protein-protein Interactions

High-density functional protein microarrays containing ~4,200 recombinant yeast proteins are examined for kinase protein-protein interactions using an affinity purified yeast kinase fusion protein containing a V5-epitope tag for read-out. Purified kinase is obtained through culture of a yeast strain optimized for high copy protein production harboring a plasmid containing a Kinase-V5 fusion construct under a GAL inducible promoter. The yeast is grown in restrictive media with a neutral carbon source for 6 hr followed by induction with 2% galactose. Next, the culture is harvested and kinase is purified using standard affinity chromatographic techniques to obtain a highly purified protein kinase for use in the assay. The purified kinase is diluted with kinase buffer to an appropriate range for the assay and the protein microarrays are blocked prior to hybridization with the protein microarray. After the hybridization, the arrays are probed with monoclonal V5 antibody to identify proteins bound by the kinase-V5 protein. Finally, the arrays are scanned using a standard microarray scanner, and data is extracted for downstream informatics analysis1,2 to determine a high confidence set of protein interactions for downstream validation in vivo.

Using protein microarrays containing nearly the entire S. cerevisiae proteome is probed for rapid unbiased interrogation of thousands of protein-protein interactions in parallel. This method can be utilized for protein-small molecule, posttranslational modification, and other assays in high-throughput.

High-density functional protein microarrays containing ~4,200 recombinant yeast proteins are examined for kinase protein-protein interactions using an ...

Development of Inhibitors of Protein-protein Interactions through REPLACE: Application to the Design and Development Non-ATP Competitive CDK Inhibitors

REPLACE is a unique strategy developed to more effectively target protein-protein interactions (PPIs). It aims to expand available drug target space by providing improved methodology for the identification of inhibitors for such binding sites and which represent the majority of potential drug targets. The main goal of this paper is to provide a methodological overview of the use and application of the REPLACE strategy which involves computational and synthetic chemistry approaches. REPLACE is exemplified through its application to the development of non-ATP competitive cyclin dependent kinases (CDK) inhibitors as anti-tumor therapeutics. CDKs are frequently deregulated in cancer and hence are considered as important targets for drug development. Inhibition of CDK2/cyclin A in S phase has been reported to promote selective apoptosis of cancer cells in a p53 independent manner through the E2F1 pathway. Targeting the protein-protein interaction at the cyclin binding groove (CBG) is an approach which will allow the specific inhibition of cell cycle over transcriptional CDKs. The CBG is recognized by a consensus sequence derived from CDK substrates and tumor suppressor proteins termed the cyclin binding motif (CBM). The CBM has previously been optimized to an octapeptide from p21Waf (HAKRRIF) and then further truncated to a pentapeptide retaining sufficient activity (RRLIF). Peptides in general are not cell permeable, are metabolically unstable and therefore the REPLACE (REplacement with Partial Ligand Alternatives through Computational Enrichment) strategy has been applied in order to generate more drug-like inhibitors. The strategy begins with the design of Fragment ligated inhibitory peptides (FLIPs) that selectively inhibit cell cycle CDK/cyclin complexes. FLIPs were generated by iteratively replacing residues of HAKRRLIF/RRLIF with fragment like small molecules (capping groups), starting from the N-terminus (Ncaps), followed by replacement on the C-terminus. These compounds are starting points for the generation of non-ATP competitive CDK inhibitors as anti-tumor therapeutics.

We describe implementation of the REPLACE strategy for targeting protein-protein interactions. REPLACE is an iterative strategy involving synthetic and computational approaches for the conversion of optimized peptidic inhibitors into drug like molecules.

REPLACE is a unique strategy developed to more effectively target protein-protein interactions (PPIs). It aims to expand available drug target space by ...

Chromatin Immunoprecipitation Assay for the Identification of Arabidopsis Protein-DNA Interactions In Vivo

Intricate gene regulatory networks orchestrate biological processes and developmental transitions in plants. Selective transcriptional activation and silencing of genes mediate the response of plants to environmental signals and developmental cues. Therefore, insights into the mechanisms that control plant gene expression are essential to gain a deep understanding of how biological processes are regulated in plants. The chromatin immunoprecipitation (ChIP) technique described here is a procedure to identify the DNA-binding sites of proteins in genes or genomic regions of the model species Arabidopsis thaliana. The interactions with DNA of proteins of interest such as transcription factors, chromatin proteins or posttranslationally modified versions of histones can be efficiently analyzed with the ChIP protocol. This method is based on the fixation of protein-DNA interactions in vivo, random fragmentation of chromatin, immunoprecipitation of protein-DNA complexes with specific antibodies, and quantification of the DNA associated with the protein of interest by PCR techniques. The use of this methodology in Arabidopsis has contributed significantly to unveil transcriptional regulatory mechanisms that control a variety of plant biological processes. This approach allowed the identification of the binding sites of the Arabidopsis chromatin protein EBS to regulatory regions of the master gene of flowering FT. The impact of this protein in the accumulation of particular histone marks in the genomic region of FT was also revealed through ChIP analysis.

Chromatin Immunoprecipitation Assay for the Identification of Arabidopsis Protein-DNA Interactions In Vivo

Chromatin immunoprecipitation is a powerful technique for the identification of DNA binding sites of Arabidopsis proteins in vivo. This procedure includes chromatin cross-linking and fragmentation, immunoprecipitation with selective antibodies against the protein of interest, and qPCR analysis of bound DNA. We describe a simple ChIP assay optimized for Arabidopsis plants.

Intricate gene regulatory networks orchestrate biological processes and developmental transitions in plants. Selective transcriptional activation and ...

Study of Protein-protein Interactions in Autophagy Research

Protein-protein interactions are important for understanding cellular signaling cascades and identifying novel pathway components and protein dynamics. The majority of cellular activities require physical interactions between proteins. To analyze and map these interactions, various experimental techniques as well as bioinformatics tools were developed. Autophagy is a cellular recycling mechanism that allows the cells to cope with different stressors, including nutrient deprivation, chemicals, and hypoxia. In order to better understand autophagy-related signaling events and to discover novel factors that regulate protein complexes in autophagy, we performed protein-protein interaction screens. Validation of these screening results requires the use of immunofluorescence and immunoprecipitation techniques. In this system, specific autophagy-related protein-protein interactions that we discovered were tested in Neuro2A (N2A) and HEK293T cell lines. Details of the technical procedures used are explained in this visualized experiment paper.

Presented here are two antibody-based protein-protein interaction research techniques: immunofluorescence and immunoprecipitation. These techniques are suitable for studying physical interactions between proteins for the discovery of novel components of cellular signaling pathways and for understanding protein dynamics.

Protein-protein interactions are important for understanding cellular signaling cascades and identifying novel pathway components and protein dynamics. ...

Advanced Confocal Microscopy Techniques to Study Protein-protein Interactions and Kinetics at DNA Lesions

Local microirradiation with lasers represents a useful tool for studies of DNA-repair-related processes in live cells. Here, we describe a methodological approach to analyzing protein kinetics at DNA lesions over time or protein-protein interactions on locally microirradiated chromatin. We also show how to recognize individual phases of the cell cycle using the Fucci cellular system to study cell-cycle-dependent protein kinetics at DNA lesions. A methodological description of the use of two UV lasers (355 nm and 405 nm) to induce different types of DNA damage is also presented. Only the cells microirradiated by the 405-nm diode laser proceeded through mitosis normally and were devoid of cyclobutane pyrimidine dimers (CPDs). We also show how microirradiated cells can be fixed at a given time point to perform immunodetection of the endogenous proteins of interest. For the DNA repair studies, we additionally describe the use of biophysical methods including FRAP (Fluorescence Recovery After Photobleaching) and FLIM (Fluorescence Lifetime Imaging Microscopy) in cells with spontaneously occurring DNA damage foci. We also show an application of FLIM-FRET (Fluorescence Resonance Energy Transfer) in experimental studies of protein-protein interactions.

Laser microirradiation is a useful tool for studies of DNA repair in living cells. A methodological approach for the use of UVA lasers to induce various DNA lesions is shown. We have optimized a method for local microirradiation that maintains the normal cell cycle; thus, irradiated cells proceed through mitosis.

Local microirradiation with lasers represents a useful tool for studies of DNA-repair-related processes in live cells. Here, we describe a methodological ...