Name: assays for the degradation of misfolded proteins in cells
Uploaded: 2016-08-28T15:00:00
Description: Watch this Scientific Journal Video about Assays for the Degradation of Misfolded Proteins in Cells at JoVE.com

We thank S. Raychaudhuri for providing the destabilized firefly luciferase mutant plasmid, and A. Glavis-bloom and N. Charan for technical assistance. This work was supported, in part, by grants from NIH (CA088868, GM060911, and CA182675).

1. Preparation of Reagent
<ol>
	<li>Prepare cell lysis buffer (50 mM Tris, pH 8.8, 100 mM NaCl, 5 mM MgCl2, 0.5% NP-40). Supplement 2 mM DTT, 1x complete protease cocktail, and 250 IU/ml benzonase before use.</li>
	<li>Prepare pellet buffer (20 mM Tris, pH 8.0, 15 mM MgCl2). Supplement 2 mM DTT, 1x complete protease cocktail and 250 IU/ml benzonase before use.</li>
	<li>Prepare 3x boiling buffer (6% SDS, 20 mM Tris, pH 8.0). Supplement 150 mM DTT before use.</li>
	<li>Prepare low-fluorescence DMEM...

<ol>
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Mechanisms that regulate the degradation of misfolded proteins are essential for maintaining the homeostasis of cellular proteins, and they likely represent valuable drug targets for treating neurodegenerative disorders and other protein-misfolding diseases. Here, assays that examine the degradation of misfolded proteins are described, using a pathogenic Atxn1 protein (Atxn1 82Q) and a nuclear localized luciferase mutant (NLS-LucDM) as examples.
To examine the degradation Atxn1 82Q, which has ...

Proteins are the most abundant macromolecules in cells, and they play an essential role in virtually all biological processes. The biological activity of most proteins requires their folding into, and maintaining, the native three-dimensional structures. Proteins with aberrant conformations not only lose their normal functions, but also frequently form soluble oligomeric species or aggregates that impair the functions of other proteins and are toxic to cells1,2. To counteract protein misfolding, cells employ both molecular chaperones, which assist unfolded or partially folded polypeptides to reach their native conformation, and degradation pathways, which eliminate misfolded proteins3. Given the complexity and stochastic nature of the folding process, protein misfolding is inevitable, and it may not be reversed in the case of mutations, biosynthetic errors, and post-translational damages1. Hence, cells ultimately rely on degradation pathways to maintain their protein quality.
The importance of cellular protein quality control (PQC) systems is underscored by the prevalence of protein-misfolding diseases, including cancer, diabetes, and many neurodegenerative disorders such as Alzheimer&#39;s disease, Parkinson&#39;s disease, amyotrophic lateral sclerosis, Huntington&#39;s disease (HD), and spinocerebellar ataxias (SCA)4,5. For example, mutations in the tumor suppressor p53 is the single most frequently genetic lesion in tumors, associated with ~50-70% of all cases6. A substantial fraction of p53 mutations are missense mutations that alter the conformation of p53, leading to the formation of aggregates7. Furthermore, proteins with expanded polyQ stretches are genetically and pathologically associated with HD and SCA. These progressive and often fatal diseases manifest when the length of the polyQ stretch in the affected proteins exceeds a certain threshold, and becomes increasingly severe as the length of the polyQ stretch prolongs8.
An attractive approach for treating these diseases is to therapeutically bolster cellular PQC systems, especially the degradation pathways. However, the pathways involved in the degradation of defective proteins, particularly those in mammalian cells, remain poorly understood. Although it has been recognized that the proteasome is critically important for the degradation of misfolded proteins, a vital issue remains undefined: how misfolded proteins are specifically recognized and targeted for degradation. Moreover, although PQC systems have been identified in cellular compartments including the cytoplasm, the endoplasmic reticulum, and the mitochondrion, the PQC systems in the nucleus remain unclear2.
Recent studies by our lab have identified a system that recognizes and degrades a variety of misfolded proteins in the nucleus of mammalian cells9. This system is comprised of the promyelocytic protein (PML), a nuclear protein and a member of the tripartite motif-containing (TRIM) protein family, and RNF4, a RING-domain containing protein. PML, and several other TRIM proteins, possess SUMO (small ubiquitin-like modifier) E3 ligase activity, which facilitates the specificity and efficiency of protein SUMOylation10. RNF4 belongs to a small group of SUMO-targeted ubiquitin ligases (STUbL), which contain one or more sumo-interacting motifs (SIMs) in addition to the RING domain that affords them the ubiquitin ligase activity11. We found that PML specifically recognizes misfolded proteins through discrete substrate recognition sites that can discern distinct features on misfolded proteins. Upon binding, PML tags misfolded proteins with poly-chains of SUMO2/3, two nearly identical mammalian SUMO proteins that can form poly-chains due to the existence of an internal SUMOylation site. SUMOylated misfolded proteins are then recognized by RNF4, which leads to their ubiquitination and proteasomal degradation. We further demonstrated that the PML-RNF4 system is important for protection against neurodegeneration, as deficiency in PML exacerbates the behavioral and neuropathological defects of a mouse model of SCA type 1 (SCA1)9.
To distinguish protein degradation from other cellular mechanisms that may regulate protein levels, the rates of protein turnover was measured9. Among the most frequently used methods to determine protein turnover are pulse chase and cycloheximide (CHX) chase. These two methods examine over time, respectively, the radioisotope-labeled proteins of interest in translation-proficient cells and total preexisting proteins of interest in translation-inhibited cells. However, a major challenge for studying pathogenic and misfolding-prone proteins is that the half-life of these proteins can be extremely long. For example, Ataxin-1, Ataxin-7, Huntingtin, &#945;-synuclein, and TDP-43 all have half-lives of more than 12 to 24 hr9,12-16. The slow turnover rates of these proteins preclude the use of the CHX chase analysis because the cells harboring these proteins may not survive prolonged translation inhibition, especially because the misfolded proteins themselves can be highly toxic to cells. The pulse-chase analysis with isotopic labeling may also be challenging for proteins that are highly aggregation-prone. Most pulse-chase assays rely on immunoprecipitation to separate the protein of interest from all the other proteins that are also radioactively labeled. This procedure normally includes lengthy immunoprecipitation and wash procedures, during which SDS-insoluble aggregates can form, making the analysis with SDS-PAGE electrophoresis inaccurate.
Here a protocol to analyze nuclear misfolded proteins with a slow turnover rate is described9. A pathogenic form of Ataxin-1 (Atxn1) that contains a stretch of 82 glutamines (Atxn1 82Q) is used for this purpose8. When expressed in cells, an enhanced green fluorescent protein (GFP) fusion of Atxn1 82Q forms microscopically visible inclusions in the nucleus (Figure 1A). Pulse chase analysis reveals that the half-life of Ataxn1 82Q is over 18 hr9. Atxn1 82Q is composed of species with misfolded conformations of different characteristics, as well as species with native conformation. It is likely that these species are degraded at different rates, and thus should be analyzed separately. Lysates from Atxn1 82Q-GFP-expressing cells are fractionated into NP-40-soluble (soluble or NS, likely representing native proteins or misfolded monomeric/oligomeric proteins) and NP-40-insoluble (NI, aggregated/misfolded) portions. The latter can be further divided into SDS-soluble (SS; likely disordered aggregates) or SDS-resistant (SR; likely amyloid fibrils) fractions (Figure 1B). NS and SS fractions can be analyzed by SDS-PAGE followed by western blotting, whereas the SR fraction can be detected by filter retardation assays followed by immunoblotting. CHX chase is combined with the detergent fractionation method, and discovered that the half-life of SS Atxn1 82Q is much shorter than that of NS Atxn1 82Q and total Atxn1 82Q (Figure 2A), indicating that the SS fraction can be readily recognized and degraded in cells9. Thus, this method provides a powerful tool to study the dynamics of misfolded proteins and to compare their degradation pattern.
We also describe a method that is suitable for high throughput screening for identifying macromolecules or small compounds that can modulate the degradation of misfolded proteins. This method is based on a conformationally destabilized mutant of firefly luciferase (LucDM)17, a model chaperone substrate. We have fused LucDM to a nuclear localization signal (NLS) to probe the PQC systems in this cellular compartment, and GFP (NLS-LucDM-GFP) for convenience of detection. NLS-LucDM-GFP forms microscopically visible nuclear aggregates in a small percentage of cells (Figure 3A). Similar to Atxn1 82Q, NLS-LucDM-GFP-but not its wild-type counterpart NLS-LucWT-GFP-is modified by SUMO2/3 and regulated by the PML-RNF4 pathway9. NLS-LucDM-GFP also forms SS and SR species, although the relative amounts of SS and SR species are minimal compared to NS, using the conditions described in protocol 3 below. To simplify the assay, we only analyze SDS soluble LucDM (including both the NS and SS fractions) by SDS-PAGE and western blot. Importantly, CHX chase assay showed that the half-life of SDS-soluble NLS-LucDM-GFP is much shorter than that of its wild-type counterpart (Figure 3B), suggesting that LucDM is a specific substrate for the system that recognizes and degrades misfolded proteins.
The degradation of LucDM-GFP causes a significant drop in overall fluorescence signal. Therefore, we have also developed a protocol for real-time detection of cellular LucDM-GFP using microplate fluorescence-based assay. Many high-throughput screen (HTS) systems are developed for drugs or genes modifying cellular aggregates and cellular viability caused by aberrant proteins18-20. However, very few HTSs are specifically designed for targeting degradation in mammalian cells. This protocol serves as a robust system for rapid and large-scale analysis of the effects of protein expression, knockdown, and drug treatment on cellular misfolded protein degradation. Using HeLa cells as example, below we describe the protocols for the analysis of these two misfolded proteins. The assays can also be applied to other cell lines, although transfection conditions and time course may need to be optimized for individual cell lines.

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		<tr height="21">
			<td height="21" style="height:21px;width:308px;">Dulbecco&#39;s Modified Eagle Medium</td>
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			<td style="width:119px;">11995-092</td>
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			<td height="21" style="height:21px;width:308px;">Fetal Bovine Serum</td>
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			<td style="width:119px;">10082147</td>
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			<td height="21" style="height:21px;width:308px;">Lipofectamine 2000</td>
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			<td style="width:119px;">11668019</td>
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			<td height="21" style="height:21px;width:308px;">NP-40</td>
			<td style="width:175px;">Sigma-Aldrich</td>
			<td style="width:119px;">NP40S-500ML</td>
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			<td height="21" style="height:21px;width:308px;">SDS</td>
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			<td height="21" style="height:21px;width:308px;">MG132</td>
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			<td height="21" style="height:21px;width:308px;">Cycloheximide</td>
			<td style="width:175px;">Sigma-Aldrich</td>
			<td style="width:119px;">C7698</td>
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			<td height="21" style="height:21px;width:308px;">Dithiothreitol (DTT)</td>
			<td style="width:175px;">Fisher Scientific</td>
			<td style="width:119px;">45000232</td>
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		</tr>
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			<td height="42" style="height:42px;width:308px;">Complete Protease Inhibitor Cocktail Tablets</td>
			<td style="width:175px;">Roche Boehringer Mannheim</td>
			<td style="width:119px;">4693159001</td>
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		</tr>
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			<td height="21" style="height:21px;width:308px;">Bio-Dot Apparatus&#160;</td>
			<td style="width:175px;">Bio-Rad</td>
			<td style="width:119px;">1706545</td>
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			<td height="21" style="height:21px;width:308px;">Living Colors GFP Monoclonal Antibody</td>
			<td style="width:175px;">Clonetech</td>
			<td style="width:119px;">632375</td>
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			<td height="21" style="height:21px;width:308px;">Anti-Actin mAb Rabbit, IgG</td>
			<td style="width:175px;">Sigma-Aldrich</td>
			<td style="width:119px;">A45060-200UL</td>
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			<td height="21" style="height:21px;width:308px;">Amino acids</td>
			<td style="width:175px;">Sigma-Aldrich</td>
			<td style="width:119px;"></td>
			<td>Amino acids are used for making low fluorecence culturing medium</td>
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			<td height="42" style="height:42px;width:308px;">Olympus IX-81&#160; Inverted Fluorescence Microscope</td>
			<td style="width:175px;">Olympus</td>
			<td style="width:119px;">IX71/IX81</td>
			<td></td>
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		<tr height="42">
			<td height="42" style="height:42px;width:308px;">96 Well Black TC Plate w/ Transluscent Clear Bottom</td>
			<td style="width:175px;">Sigma-Greiner</td>
			<td style="width:119px;">89135-048</td>
			<td></td>
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		<tr height="42">
			<td height="42" style="height:42px;width:308px;">Fluorescence Bottom Plate Reader Infinite 200&#174; PRO</td>
			<td style="width:175px;">TECAN</td>
			<td style="width:119px;">Infinite 200&#174; PRO</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;width:308px;">Cellulose acetate membrane 0.2 &#181;m</td>
			<td style="width:175px;">Sterlitech</td>
			<td style="width:119px;">CA023001</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:308px;">Prism 5</td>
			<td style="width:175px;">GraphPad</td>
			<td style="width:119px;"></td>
			<td>Statistical analysis software</td>
		</tr>
	</tbody>
</table>

assays for the degradation of misfolded proteins in cells

Protein misfolding and aggregation are associated with various neurodegenerative diseases. Cellular mechanisms that recognize and degrade misfolded proteins may serve as potential therapeutic targets. To distinguish degradation of misfolding-prone proteins from other mechanisms that regulate their levels, one important method is to measure protein half-life in cells. However, this can be challenging because misfolding-prone proteins may exist in different forms, including the native form and misfolded forms of distinct characteristics. Here we describe assays to examine the half-life of misfolded proteins in mammalian cells using a highly aggregation-prone protein, Ataxin-1 with an extended polyglutamine (polyQ) stretch, and a conformationally unstable luciferase mutant as models. Cycloheximide chase is combined with cell fractionation to examine the turnover rate of misfolding-prone proteins in various cellular fractions. We further depict a fluorescence-based assay using an enhanced green fluorescence protein (EGFP)-fusion of the luciferase mutant, which can be adapted for high throughput screening on a microplate-reader.

In a steady state analysis, microscopically visible Atxn1 82Q-GFP nuclear aggregates can be observed in 30 - 50% of HeLa cells 20 hr after transfection (Figure 1A). Western blot analysis of NS and SS fractions using anti-GFP antibody shows a distinct band of Atxn1 82Q-GFP between 100 kDa and 150 kDa markers, corresponding to the protein&#39;s molecular weight (Figure 1B). Atxn1 82Q-GFP in the SR fraction can be detected either by filter retardation assay,...

Watch this Scientific Journal Video about Assays for the Degradation of Misfolded Proteins in Cells at JoVE.com

Assays for the Degradation of Misfolded Proteins in Cells

This report describes protocols for measuring degradation rates of misfolded proteins by either western blot or fluorescence-based assays. The methods can be applied to analysis of other misfolded proteins and for high throughput screening.

Protein misfolding and aggregation are associated with various neurodegenerative diseases. Cellular mechanisms that recognize and degrade misfolded ...

The overall goal of this assay is to measure the cellular degradation rates of misfolded proteins. Misfolded proteins are associated with many neurodegenerative diseases. This assay helps to investigate the cellular protein quality control systems that are protectors against aberrant proteins.Misfolding prone proteins may exist in different forms in cells. By combining cell fractionation with the cyclohexane method chase, the main advantage of this technique is to specifically measure the half life of the misfolded species. We use two model proteins as examples.A highly aggregation prone polyglutamate protein, Atxn1 82Q and a conformationally unstable nuclear luciferase mutant. The protocol can also be applied to other misfolded protein measurements. To carry out a degradation assay for Atxn1 82Q GFP, plate approximately 3x10^5 HeLa cells in 35 millimeter plates with DMEM medium and 10%FBS.Incubate the cells overnight so that they reach 40 to 60%confluence at the time of transfection. Four to five hours after transfecting the cells with Atxn1 82Q GFP PRK5, under a florescence microscope, use an excitation wavelength of 450 to 490 nanometers to examine the live cells for GFP expression in the nuclei, characterized by the presence of diffused as well as small speckles of GFP signals. Just before cycloheximide treatment, harvest one plate of cells by removing the medium and using three milliliters of ice cold PBS to wash the cells twice.Then snap freeze the plate on dry ice. For the remaining plates of cells, remove the medium by vacuum aspiration and add two milliliters of fresh DMEM containing 50 micrograms per milliliter of cycloheximide. Also add 10 micromolar of the proteasome inhibitor, MG132 to one plate.Incubate treated cells for four, eight, 12 and 16 hours before freezing on dry ice. After harvesting the MG132 treated cells at 16 hours, scrape the frozen cells from all the plates into 150 microliter of ice cold cell lysis buffer and incubate on ice for 30 minutes. Centrifuge the lysates in a benchtop centrifuge at 17, 000x g and four degrees Celsius for 15 minutes.Then transfer the supernatant which contains MP40 soluble proteins to another tube. Rinse the pellets by gently adding approximately 200 microliters of 1x PBS to the tubes without disturbing the pellets. Carefully remove the PBS by aspiration or pipette, resuspend the pellets in 150 microliters of ice cold pellet buffer and then incubate them on ice for 15 to 30 minutes.Next add 75 microliters of 3x boiling buffer into MP40 soluble fractions and MP40 insoluble fractions resuspended from the pellets. Then heat the samples at 95 degrees Celsius on a heat block for five minutes. Add SDS gel loading buffer to an aliquot of boiled MP40 soluble and insoluble fractions.Load equal volumes of samples collected from all time points onto an SDS-PAGE gel. Detect MP40 soluble and SDS soluble Atxn1 82Q GFP by Western blot using anti GFP antibody and enhanced chemiluminescence. To examine SDS resistant Atxn1 82Q from the pellet fraction with a filter retardation assay, set up a dot plot apparatus holding a 0.2 micron cellulose acetate membrane.Then load 80 to 120 microliters of boiled MP40 insoluble samples into each well of the dot blot apparatus. After filtering the samples through the membrane by vacuum, detect Atxn1 82Q GFP aggregates stuck on the membrane by anti GFP immunoblotting. It is critical to use a filter retardation assay to examine the levels of SDS resistant form over time.This is because SDS soluble form may transform into SDS resistant form rather than being degraded. In this example, SDS resistant form is minimal and remains a similar level over the chase experiment. To carry out a degradation assay, after an overnight transfection of HeLa cells with NLS-luciferase-GFP, examine the live cells under an inverted fluorescent microscope for GFP expression with an excitation wavelength of 450 to 490 nanometers.Just before cycloheximide treatment, harvest one plate of cells by removing the medium and use three milliliters of ice cold PBS to wash the cells twice. Then snap freeze the plate on dry ice. For the remaining plates, after removing the medium, add two milliliters of fresh DMEM containing 50 micrograms per milliliter of cycloheximide and add 10 micromolar of the proteasome inhibitor MG132 to one plate.Treat the cells for 1.5, three, 4.5 and six hours before harvesting as just described, harvesting the MG132 treated cells at six hours. After harvesting the last time point of cells, scrape each plate into 150 microliters of ice cold cell lysis buffer and incubate on ice for 30 minutes. Incubate the samples on heat block at 95 degrees Celsius for five minutes, before carrying out SDS-PAGE and Western blotting according to the text protocol.Add SDS gel loading buffer to whole cell lysates for a final concentration of 2%SDS and 50 millimolar DTT. After seeding and transfecting HeLa cells in 96 well plates according to the text protocol, 20 to 24 hours after transfection, examine the cells for GFP expression. Remove the medium, add approximately 200 microliters of 1x PBS to each well and then aspirate it to remove the residual DMEM.Add 60 microliters of low fluorescence DMEM with 5%FBS and 50 micrograms per milliliter of cycloheximide and add 10 micromolar MG132 to one set of samples. With a fluorescence plate reader, measure the GFP signal, reading the plates every hour for up to eight to 10 hours. Export the data and carry out statistical analysis according to the text protocol.In a steady state analysis, microscopically visible Atxn1 82Q GFP nuclear aggregates can be observed in 30 to 50%of HeLa cells 20 hours after transfection. As seen here by Western blot, the half life of Atxn1 82Q GFP in the SDS soluble fraction is around five hours. In contrast to a slight or no decrease in Atxn1 82Q GFP in the MP40 soluble fraction over 16 hours.The degradation of Atxn1 82Q GFP is partially inhibited by treating cells with MG132. These images illustrate that 20 hours after transfection, NLS-luciferase DM mutant GFP aggregates become microscopically visible in five to 15%of HeLa cells. Western blots revealed that the half life of SDS soluble NLS-luciferase DM mutant GFP is two to three hours.In addition, the florescence intensity of transfected cells also decreases over time upon cycloheximide treatment. At approximately six hours after cycloheximide treatment, soluble NLS-luciferase DM mutant GFP is largely degraded suggesting that the remaining fluorescence is generated from aggregated species resistant to degradation. These florescence images confirm that aggregated, but not diffused GFP, remains in cells nine hours after cycloheximide treatment.Well the immunblotting based assay takes a few days to accomplish. The protein degradation rates can be obtained immediately after the cycloheximide chase using a fluorescence based assay. Both immunoblotting and fluorescence based assays can be applied to studies of other misfolded protein.The latter is also suitable for high throughput screening for identifying macromolecules or small compounds that can modulate the degradation of misfolded proteins. It is important to remember that for some misfolded proteins such Atxn1 82Q, the half life can only be measured through cell fractionation and immunoblotting and this is because the misfolded species that degraded quickly are only a small portion of the overall Atxn1 82Q expression. To uncover the mechanism of a protein quality controls, protein degradation assays need to be coupled with analysis of the steady state levels of aggregates as well as that they are partitions in different cell fractions.Other experiments like in vitro protein folding and aggregation assays can also be performed. After watching this video, you should have a good understanding of how to measure the degradation rates of misfolding proteins using different strategies. Thanks for watching and good luck with your experiments.

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Research

JoVE Journal

Biology

Crystallization of Membrane Proteins in Lipidic Mesophases

Membrane proteins perform critical functions in living cells related to signal transduction, transport and energy transformations, and, as such, are implicated in a multitude of malfunctions and diseases. However, a structural and functional understanding of membrane proteins is strongly lagging behind that of their soluble partners, mainly, due to difficulties associated with their solubilization and generation of diffraction quality crystals. Crystallization in lipidic mesophases (also known as in meso or LCP crystallization) is a promising technique which was successfully applied to obtain high resolution structures of microbial rhodopsins, photosynthetic proteins, outer membrane beta barrels and G protein-coupled receptors. In meso crystallization takes advantage of a native-like membrane environment and typically produces crystals with lower solvent content and better ordering as compared to traditional crystallization from detergent solutions. The method is not difficult, but requires an understanding of lipid phase behavior and practice in handling viscous mesophase materials. Here we demonstrate a simple and efficient way of making LCP and reconstituting a membrane protein in the lipid bilayer of LCP using a syringe mixer, followed by dispensing nanoliter portions of LCP into an assay or crystallization plate, conducting pre-crystallization assays and harvesting crystals from the LCP matrix. These protocols provide a basic guide for approaching in meso crystallization trials; however, as with any crystallization experiment, extensive screening and optimization are required, and a successful outcome is not necessarily guaranteed.

The protocols describe the essential steps for obtaining diffraction quality crystals of a membrane protein starting from reconstitution of the protein in a lipidic cubic phase (LCP), finding initial conditions with LCP-FRAP pre-crystallization assays, setting up LCP crystallization trials and harvesting crystals.

Membrane proteins perform critical functions in living cells related to signal transduction, transport and energy transformations, and, as such, are ...

Photobleaching Assays (FRAP &amp; FLIP) to Measure Chromatin Protein Dynamics in Living Embryonic Stem Cells

Fluorescence Recovery After Photobleaching (FRAP) and Fluorescence Loss In Photobleaching (FLIP) enable the study of protein dynamics in living cells with good spatial and temporal resolution. Here we describe how to perform FRAP and FLIP assays of chromatin proteins, including H1 and HP1, in mouse embryonic stem (ES) cells. In a FRAP experiment, cells are transfected, either transiently or stably, with a protein of interest fused with the green fluorescent protein (GFP) or derivatives thereof (YFP, CFP, Cherry, etc.). In the transfected, fluorescing cells, an intense focused laser beam bleaches a relatively small region of interest (ROI). The laser wavelength is selected according to the fluorescent protein used for fusion. The laser light irreversibly bleaches the fluorescent signal of molecules in the ROI and, immediately following bleaching, the recovery of the fluorescent signal in the bleached area - mediated by the replacement of the bleached molecules with the unbleached molecules - is monitored using time lapse imaging. The generated fluorescence recovery curves provide information on the protein's mobility. If the fluorescent molecules are immobile, no fluorescence recovery will be observed. In a complementary approach, Fluorescence Loss in Photobleaching (FLIP), the laser beam bleaches the same spot repeatedly and the signal intensity is measured elsewhere in the fluorescing cell. FLIP experiments therefore measure signal decay rather than fluorescence recovery and are useful to determine protein mobility as well as protein shuttling between cellular compartments. Transient binding is a common property of chromatin-associated proteins. Although the major fraction of each chromatin protein is bound to chromatin at any given moment at steady state, the binding is transient and most chromatin proteins have a high turnover on chromatin, with a residence time in the order of seconds. These properties are crucial for generating high plasticity in genome expression1. Photobleaching experiments are therefore particularly useful to determine chromatin plasticity using GFP-fusion versions of chromatin structural proteins, especially in ES cells, where the dynamic exchange of chromatin proteins (including heterochromatin protein 1 (HP1), linker histone H1 and core histones) is higher than in differentiated cells2,3.

Photobleaching Assays FRAP &amp; FLIP to Measure Chromatin Protein Dynamics in Living Embryonic Stem Cells

We describe photobleaching methods including Fluorescence Recovery After Photobleaching (FRAP) and Fluorescence Loss In Photobleaching (FLIP) to monitor chromatin protein dynamics in embryonic stem (ES) cells. Chromatin protein dynamics, which is considered to be one of the means to study chromatin plasticity, is enhanced in pluripotent cells.

Fluorescence Recovery After Photobleaching (FRAP) and Fluorescence Loss In Photobleaching (FLIP) enable the study of protein dynamics in living cells with ...

Viability Assays for Cells in Culture

Manual cell counts on a microscope are a sensitive means of assessing cellular viability but are time-consuming and therefore expensive. Computerized viability assays are expensive in terms of equipment but can be faster and more objective than manual cell counts. The present report describes the use of three such viability assays. Two of these assays are infrared and one is luminescent. Both infrared assays rely on a 16&#160;bit Odyssey Imager. One infrared assay uses the DRAQ5 stain for nuclei combined with the Sapphire stain for cytosol and is visualized in the 700 nm channel. The other infrared assay, an In-Cell Western, uses antibodies against cytoskeletal proteins (&#945;-tubulin or microtubule associated protein 2) and labels them in the 800 nm channel. The third viability assay is a commonly used luminescent assay for ATP, but we use a quarter of the recommended volume to save on cost. These measurements are all linear and correlate with the number of cells plated, but vary in sensitivity. All three assays circumvent time-consuming microscopy and sample the entire well, thereby reducing sampling error. Finally, all of the assays can easily be completed within one day of the end of the experiment, allowing greater numbers of experiments to be performed within short timeframes. However, they all rely on the assumption that cell numbers remain in proportion to signal strength after treatments, an assumption that is sometimes not met, especially for cellular ATP. Furthermore, if cells increase or decrease in size after treatment, this might affect signal strength without affecting cell number. We conclude that all viability assays, including manual counts, suffer from a number of caveats, but that computerized viability assays are well worth the initial investment. Using all three assays together yields a comprehensive view of cellular structure and function.

Therapeutic compounds are often first examined in vitro with viability assays. Blind cell counts by a human observer can be highly sensitive to small changes in cell number but do not assess function. Computerized viability assays, as described here, can assess both structure and function in an objective manner.

Manual cell counts on a microscope are a sensitive means of assessing cellular viability but are time-consuming and therefore expensive. Computerized ...

The Cell-based L-Glutathione Protection Assays to Study Endocytosis and Recycling of Plasma Membrane Proteins

Membrane trafficking involves transport of proteins from the plasma membrane to the cell interior (i.e. endocytosis) followed by trafficking to lysosomes for degradation or to the plasma membrane for recycling. The cell based L-glutathione protection assays can be used to study endocytosis and recycling of protein receptors, channels, transporters, and adhesion molecules localized at the cell surface. The endocytic assay requires labeling of cell surface proteins with a cell membrane impermeable biotin containing a disulfide bond and the N-hydroxysuccinimide (NHS) ester at 4 &#186;C -&#160;a temperature at which membrane trafficking does not occur. Endocytosis of biotinylated plasma membrane proteins is induced by incubation at 37 &#186;C. Next, the temperature is decreased again to 4 &#186;C to stop endocytic trafficking and the disulfide bond in biotin covalently attached to proteins that have remained at the plasma membrane is reduced with L-glutathione. At this point, only proteins that were endocytosed remain protected from L-glutathione and thus remain biotinylated. After cell lysis, biotinylated proteins are isolated with streptavidin agarose, eluted from agarose, and the biotinylated protein of interest is detected by western blotting. During the recycling assay, after biotinylation cells are incubated at 37 &#176;C to load endocytic vesicles with biotinylated proteins and the disulfide bond in biotin covalently attached to proteins remaining at the plasma membrane is reduced with L-glutathione at 4 &#186;C as in the endocytic assay. Next, cells are incubated again at 37 &#176;C to allow biotinylated proteins from endocytic vesicles to recycle to the plasma membrane. Cells are then incubated at 4 &#186;C, and the disulfide bond in biotin attached to proteins that recycled to the plasma membranes is reduced with L-glutathione. The biotinylated proteins protected from L-glutathione are those that did not recycle to the plasma membrane.

Membrane trafficking involves transport of proteins from the plasma membrane to the cell interior (i.e. endocytosis) followed by trafficking to lysosomes for degradation or to the plasma membrane for recycling. Methods described in this article are designed to study endocytosis and recycling of plasma membrane proteins.

Membrane trafficking involves transport of proteins from the plasma membrane to the cell interior (i.e. endocytosis) followed by trafficking to lysosomes ...

Reconstitution Of &#946;-catenin Degradation In Xenopus Egg Extract

Xenopus laevis egg extract is a well-characterized, robust system for studying the biochemistry of diverse cellular processes. Xenopus egg extract has been used to study protein turnover in many cellular contexts, including the cell cycle and signal transduction pathways1-3. Herein, a method is described for isolating Xenopus egg extract that has been optimized to promote the degradation of the critical Wnt pathway component, &beta;-catenin. Two different methods are described to assess &beta;-catenin protein degradation in Xenopus egg extract. One method is visually informative ([35S]-radiolabeled proteins), while the other is more readily scaled for high-throughput assays (firefly luciferase-tagged fusion proteins). The techniques described can be used to, but are not limited to, assess &beta;-catenin protein turnover and identify molecular components contributing to its turnover. Additionally, the ability to purify large volumes of homogenous Xenopus egg extract combined with the quantitative and facile readout of luciferase-tagged proteins allows this system to be easily adapted for high-throughput screening for modulators of &beta;-catenin degradation.

Reconstitution Of &amp;#946;-catenin Degradation In Xenopus Egg Extract

A method is described for analyzing protein degradation using radiolabeled and luciferase-fusion proteins in Xenopus egg extract and its adaptation for high-throughput screening for small molecule modulators of protein degradation.

Xenopus laevis egg extract is a well-characterized, robust system for studying the biochemistry of diverse cellular processes. Xenopus egg extract has ...

Reporter-based Growth Assay for Systematic Analysis of Protein Degradation

Protein degradation by the ubiquitin-proteasome system (UPS) is a major regulatory mechanism for protein homeostasis in all eukaryotes. The standard approach to determining intracellular protein degradation relies on biochemical assays for following the kinetics of protein decline. Such methods are often laborious and time consuming and therefore not amenable to experiments aimed at assessing multiple substrates and degradation conditions. As an alternative, cell growth-based assays have been developed, that are, in their conventional format, end-point assays that cannot quantitatively determine relative changes in protein levels.
Here we describe a method that faithfully determines changes in protein degradation rates by coupling them to yeast cell-growth kinetics. The method is based on an established selection system where uracil auxotrophy of URA3-deleted yeast cells is rescued by an exogenously expressed reporter protein, comprised of a fusion between the essential URA3 gene and a degradation determinant (degron). The reporter protein is designed so that its synthesis rate is constant whilst its degradation rate is determined by the degron. As cell growth in uracil-deficient medium is proportional to the relative levels of Ura3, growth kinetics are entirely dependent on the reporter protein degradation.
This method accurately measures changes in intracellular protein degradation kinetics. It was applied to: (a) Assessing the relative contribution of known ubiquitin-conjugating factors to proteolysis (b) E2 conjugating enzyme structure-function analyses (c) Identification and characterization of novel degrons. Application of the degron-URA3-based system transcends the protein degradation field, as it can also be adapted to monitoring changes of protein levels associated with functions of other cellular pathways.

Here we describe a robust biological assay for quantifying the relative rate of proteolysis by the ubiquitin-proteasome system. The assay readout is yeast growth rate in liquid culture, which is dependent on the cellular levels of a reporter protein comprising a degradation signal fused to an essential metabolic marker.

Protein degradation by the ubiquitin-proteasome system (UPS) is a major regulatory mechanism for protein homeostasis in all eukaryotes. The standard ...

Comparing the Affinity of GTPase-binding Proteins using Competition Assays

In this protocol we demonstrate a method for comparing the competition between GTPase-binding proteins. Such an approach is important for determining the binding capabilities of GTPases for two reasons: The fact that all interactions involve the same face of the GTPases means that binding events must be considered in the context of competitors, and the fact that the bound nucleotide must also be controlled means that conventional approaches such as immunoprecipitation are unsuitable for GTPase biochemistry. The assay relies on the use of purified proteins. Purified Rac1 immobilized on beads is used as the bait protein, and can be loaded with GDP, a non-hydrolyzable version of GTP or left nucleotide free, so that the signaling stage to be investigated can be controlled. The binding proteins to be investigated are purified from mammalian cells, to allow correct folding, by means of a GFP tag. Use of the same tag on both proteins is important because not only does it allow rapid purification and elution, but also allows detection of both competitors with the same antibody during elution. This means that the relative amounts of the two bound proteins can be determined accurately.

This protocol compares the relative affinities of binding partners for Rho-family GTPases, including Rac1. In vivo, Rac1-binding proteins compete for a single binding interface, the conformation of which is dictated by a bound nucleotide. The nucleotide is both important and difficult to control experimentally, due to the high hydrolysis rate.

In this protocol we demonstrate a method for comparing the competition between GTPase-binding proteins. Such an approach is important for determining the ...

High Yield Expression of Recombinant Human Proteins with the Transient Transfection of HEK293 Cells in Suspension

The art of producing recombinant proteins with complex post-translational modifications represents a major challenge for studies of structure and function. The rapid establishment and high recovery from transiently-transfected mammalian cell lines addresses this barrier and is an effective means of expressing proteins that are naturally channeled through the ER and Golgi-mediated secretory pathway. Here is one protocol for protein expression using the human HEK293F and HEK293S cell lines transfected with a mammalian expression vector designed for high protein yields. The applicability of this system is demonstrated using three representative glycoproteins that expressed with yields between 95-120 mg of purified protein recovered per liter of culture. These proteins are the human Fc&#947;RIIIa and the rat &#945;2-6 sialyltransferase, ST6GalI, both expressed with an N-terminal GFP fusion, as well as the unmodified human immunoglobulin G1 Fc. This robust system utilizes a serum-free medium that is adaptable for expression of isotopically enriched proteins and carbohydrates for structural studies using mass spectrometry and nuclear magnetic resonance spectroscopy. Furthermore, the composition of the N-glycan can be tuned by adding a small molecule to prevent certain glycan modifications in a manner that does not reduce yield.

Laboratory-scale production of eukaryotic proteins with appropriate post-translational modification represents a significant barrier. Here is a robust protocol with rapid establishment and turnaround for protein expression using a mammalian expression system. This system supports selective amino acid, selective labeling of proteins and small molecule modulators of glycan composition.

The art of producing recombinant proteins with complex post-translational modifications represents a major challenge for studies of structure and ...

Nephrotoxin Microinjection in Zebrafish to Model Acute Kidney Injury

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid decline in renal function and development of the clinical syndrome known as acute kidney injury (AKI). Pharmacological agents used to treat medical circumstances ranging from bacterial infection to cancer, when administered individually or in combination with other drugs, can initiate AKI. Zebrafish are a useful animal model to study the chemical effects on renal function in vivo, as they form an embryonic kidney comprised of nephron functional units that are conserved with higher vertebrates, including humans. Further, zebrafish can be utilized to perform genetic and chemical screens, which provide opportunities to elucidate the cellular and molecular facets of AKI and develop therapeutic strategies such as the identification of nephroprotective molecules. Here, we demonstrate how microinjection into the zebrafish embryo can be utilized as a paradigm for nephrotoxin studies.

Renal injuries incurred from nephrotoxins, which include drugs ranging from antibiotics to chemotherapeutics, can result in complex disorders whose pathogenesis remains incompletely understood. This protocol demonstrates how zebrafish can be used for disease modeling of these conditions, which can be applied to the identification of renoprotective measures.

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid ...

A Graphical User Interface for Software-assisted Tracking of Protein Concentration in Dynamic Cellular Protrusions

Filopodia are dynamic, finger-like cellular protrusions associated with migration and cell-cell communication. In order to better understand the complex signaling mechanisms underlying filopodial initiation, elongation and subsequent stabilization or retraction, it is crucial to determine the spatio-temporal protein activity in these dynamic structures. To analyze protein function in filopodia, we recently developed a semi-automated tracking algorithm that adapts to filopodial shape-changes, thus allowing parallel analysis of protrusion dynamics and relative protein concentration along the whole filopodial length. Here, we present a detailed step-by-step protocol for optimized cell handling, image acquisition and software analysis. We further provide instructions for the use of optional features during image analysis and data representation, as well as troubleshooting guidelines for all critical steps along the way. Finally, we also include a comparison of the described image analysis software with other programs available for filopodia quantification. Together, the presented protocol provides a framework for accurate analysis of protein dynamics in filopodial protrusions using image analysis software.

We present a software solution for semi-automated tracking of relative protein concentration along the length of dynamic cellular protrusions.

Filopodia are dynamic, finger-like cellular protrusions associated with migration and cell-cell communication. In order to better understand the complex ...