Name: purification and reconstitution of trpv1 for spectroscopic analysis
Uploaded: 2018-07-03T13:30:00
Description: Watch this Scientific Journal Video about Purification and Reconstitution of TRPV1 for Spectroscopic Analysis at JoVE.com

We are very grateful to Dr. H. Mchaourab for providing access to the EPR and DEER spectrometers and Dr. T. Rosenbaum for providing the full-length cysteine-less TRPV1 plasmid.

1. TRPV1 Mutagenesis
Note: A minimal TRPV1 construct for spectroscopic analysis26 was built from the full-length cysteine-less channel TRPV127 using the polymerase chain reaction (PCR) method (Figure 1). This cysteine-less minimal TRPV1 construct (referred to as eTRPV1 hereafter) consists of residues 110 - 603 and 627 - 764. eTRPV1 was cloned in pMO (a pcDNA3.1-based vector) for functional analysis and in a recombinant...

Current technologies for expression and purification of mammalian membrane proteins have made it possible to obtain sufficient amounts of protein for spectroscopic studies14,15,16,42. Here, we have adapted these technologies to express, purify, reconstitute, and perform spectroscopic analyses in TRPV1.
Among the critical steps in the protocol, below are the ones that...

With recent advances in single-particle cryo-electron microscopy (cryo-EM), mammalian ion channel structures have been obtained at an extraordinary rate. Particularly, structural studies of polymodal ion channels, such as the transient receptor potential vanilloid 1 (TRPV1), have provided further understanding of its activation mechanisms1,2,3,4,5. However, dynamic information about ion channels embedded in a membrane environment is required to understand their polymodal gating and drug-binding mechanisms.
Electron paramagnetic resonance (EPR) and double electron-electron resonance (DEER) spectroscopies have provided some of the most definitive mechanistic models for ion channels6,7,8,9,10,11,12,13. These approaches have been mainly restricted to the examination of prokaryotic and archeal ion channels that yield a large amount of detergent-purified proteins when overexpressed in bacteria. With the development of eukaryotic membrane proteins production in insect and mammalian cells for functional and structural characterization14,15,16, it is now possible to obtain biochemical amounts of detergent-purified proteins for spectroscopic studies.
The EPR and DEER signals arise from a&#160;paramagneticspin label (SL) (i.e., methanethiosulfonate) attached to a single-cysteine residue in the protein. The spin-labels report three types of structural information: motion, accessibilities, and distances. This information allows determining whether residues are buried within the protein or are exposed to the membrane or aqueous environment in the apo and ligand-bound states13,17,18,19. In the context of a high-resolution structure (when available), the EPR and DEER data provide a collection of constraints for deriving dynamic models in their native environment while monitoring reversible gating transition (i.e., closed, open, sensitized, and desensitized). Moreover, flexible regions that might be difficult to determine by X-ray crystallography or cryo-EM could be obtained by using these environmental data sets to assign secondary structures as well as location within the protein20. Cryo-EM structures obtained in lipid nanodiscs provided valuable information about the gating of ion channels3,21,22,23,24,25; however, spectroscopic approaches could provide dynamic information from conformational states (e.g., thermal changes) that might be difficult to determine using cryo-EM.
Many difficulties must be overcome to implement EPR and DEER, including lack of protein function when removing all cysteine residues (especially abundant in mammalian channels), low protein yield, protein instability during purification and after spin labeling, and protein aggregation in detergent or liposomes. Here, we have designed protocols to overcome these critical barriers and have obtained DEER and EPR spectra information for a mammalian sensory receptor. The purpose here is to describe methodologies for the expression, purification, labeling, and reconstitution of a functional minimal cysteine-less rat TRPV1 (eTRPV1) construct for spectroscopic analyses. This methodology is appropriate for those membrane proteins that keep their function despite the removal of cysteine residues or that contain cysteine forming disulfide-bonds. This collection of protocols could be adapted for the spectroscopic analysis of other mammalian ion channels.

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			<td height="21" style="height:21px;width:339px;">pFastBac1 Expression Vector</td>
			<td style="width:292px;">Invitrogen Life Technologies</td>
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		<tr height="21">
			<td height="21" style="height:21px;width:339px;">DH10Bac Competent Cells</td>
			<td style="width:292px;">Invitrogen Life Technologies</td>
			<td style="width:251px;">10361-012</td>
			<td style="width:167px;"></td>
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			<td>Drummond Scientific</td>
			<td>3-000-204</td>
			<td></td>
		</tr>
	</tbody>
</table>

purification and reconstitution of trpv1 for spectroscopic analysis

Polymodal ion channels transduce multiple stimuli of different natures into allosteric changes; these dynamic conformations are challenging to determine and remain largely unknown. With recent advances in single-particle cryo-electron microscopy (cryo-EM) shedding light on the structural features of agonist binding sites and the activation mechanism of several ion channels, the stage is set for an in-depth dynamic analysis of their gating mechanisms using spectroscopic approaches. Spectroscopic techniques such as electron paramagnetic resonance (EPR) and double electron-electron resonance (DEER) have been mainly restricted to the study of prokaryotic ion channels that can be purified in large quantities. The requirement for large amounts of functional and stable membrane proteins has hampered the study of mammalian ion channels using these approaches. EPR and DEER offer many advantages, including determination of the structure and dynamic changes of mobile protein regions, albeit at low resolution, that might be difficult to obtain by X-ray crystallography or cryo-EM, and monitoring reversible gating transition (i.e., closed, open, sensitized, and desensitized). Here, we provide protocols for obtaining milligrams of functional detergent-solubilized transient receptor potential cation channel subfamily V member 1 (TRPV1) that can be labeled for EPR and DEER spectroscopy.

Functional Characterization of the Minimal Cysteine-less TRPV1 Construct (eTRPV1) and Single-cysteine Mutants
The first step toward spectroscopic studies is to engineer and characterize cysteine-less protein constructs (Figure 2A) that are functional and yield biochemical amounts of proteins. eTRPV1 is functional as determined by Ca2+ imaging and TEVC (Figure 2B...

Watch this Scientific Journal Video about Purification and Reconstitution of TRPV1 for Spectroscopic Analysis at JoVE.com

Purification and Reconstitution of TRPV1 for Spectroscopic Analysis

This article describes specific methods to obtain biochemical quantities of detergent-solubilized TRPV1 for spectroscopic analysis. The combined protocols provide biochemical and biophysical tools that can be adapted to facilitate structural and functional studies for mammalian ion channels in a membrane-controlled environment.

Polymodal ion channels transduce multiple stimuli of different natures into allosteric changes; these dynamic conformations are challenging to determine ...

Here in the lab, we think that using a spectroscopic approach can help answering key questions in the transient receptor potential ion channel family. Such as determining the dynamic informational changes associated with polymodal activation. The main advantage of this protocol is that allow milligram of pure and functional remaining ion channel for electron paramagnetic resonance and double electron electron resonance epitroscopy.This method can provide insight into the conformational changes compatible with thermal and light independent gating of 3B1. It can also be applied to other trip channels, like 3B2, for example. Begin this procedure with design mutagenesis primers with online tools.Prepare the PCR reaction as described in the text protocol and spin the mixture. Place the tubes in the thermocycler and perform PCR mutagenesis using the PCR parameters listed in the text protocol. Then add two microliters of DPN1 enzyme to the reaction and mix.Incubate the reactions at 37 degrees Celsius for 30 minutes. Next, perform the transformation by adding 75 microliters of thawed E.Coli competent cells and 10 microliters of DPN1 treated PCR mixture to a pre-cooled 14 milliliter sterile tube. After mixing gently, incubate reaction on ice for 30 minutes.Maintain the tube at 42 degrees Celsius for 45 seconds and then immediately place it on ice for two minutes. Plate the mixture on an LB agar plate containing carbenicillin, and incubate overnight at 37 degrees Celsius. After harvesting single colonies and preparing overnights as described in the text protocol, spin down overnight cultures at 8000 times G for 10 minutes.Then, extract the DNA following the instructions of commercial available mini preparation kits. To generate the recombinant bacmid, transform 100 microliters of DH10bac E.Coli competent cells with 7.5 nanograms of the recombinant donor vector. Vectors contain the functional minimal cysteine-less rat TRPV1, abbreviated eTRPV1, and or single cysteine mutants.After seeding and incubating cells as described in the text protocol, select white colonies that are two millimeters in diameter since blue colonies lack the insert of interest. Use a stereo microscope to verify that white colonies do not contain any blue spots. Harvest at least three single colonies and transfer them into a 14 milliliter steril tube containing four milliliters supplemented LB media.Incubate the cells overnight at 37 degrees Celsius in 250 RPM. Take 1.5 milliliters of the overnight cultures and isolate recombinant bacmid DNA. To generate the recombinant baculovirus, plate one million SF9 cells in two millimeters of insect cell media.Allow the cells to attach for at least 45 minutes. After washing once with one milliliter of fresh medium, remove the wash medium and add DNA transfection reagent mixture to the cells. Incubate the cells at 27 degrees Celsius for five hours without agitation.Following incubation, remove the transfection mixture and replace with two milliliters of insect cell media containing 0.5%fetal bovine serum. Then, incubate the cells at 27 degrees Celsius for five days. Transfer the cell culture media into a 15 milliliter centrifuge tube.Isolate the first passage of viral stock by spinning the tube at 8000 times G for 15 minutes at 4 degrees Celsius. Transfer the supernatant to a clean 15 milliliter centrifuge tube and immediately store it at four degrees Celsius. Of the second generation of viral stock, pipette a 25 milliliter suspension culture of insect cell media at one million SF9 cells per milliliter containing 2%FBS into a 125 milliliter flask.Add 25 microliters of the P1 virus to the 25 milliliter culture. Then, incubate the culture at 27 degrees Celsius for 72 hours. To harvest the P2 viral stock transfer the culture to a new 50 milliliter tube and centrifuge it at 8000 times G for 15 minutes at four degrees Celsius.Transfer the supernatant to a new 50 milliliter tube and immediately store it at four degrees Celsius. Add one milliliter of P2 virus stock to the one liter culture and incubate it at 27 degrees Celsius for 72 hours. Harvest the one liter insect cell suspension culture by spinning at 4500 G for 20 minutes at four degrees Celsius.Weigh the cell pellet. It is expected to be around six to nine grams. Resuspend the cell pellet in 25 milliliters of ice cold buffer A in the presence of protease inhibitors.Disaggregate cell clumps to obtain a homogenous suspension and rotate at four degrees Celsius for 20 minutes. Break the cells using a manual or a high-pressure homogenizer. After removing the cell debris by centrifugation at 8000 times G for 20 minutes a four degrees Celsius, collect the supernatant and centrifuge again at 100, 000 times G for 30 minutes at four degrees Celsius.Following centrifugation, discard the supernatant and suspend the membrane pellet in a total volume of 20 milliliters of ice cold buffer B supplemented with protease inhibitors. Aliquot 20 milliliters of membrane suspension to 50 milliliter tubes. To perform the protein purification add three milliliters of DDM per tube.Rotate the tubes for two hours at four degrees Celsius. After centrifuging the sample at 100, 000 times G for 30 minutes at four degrees Celsius, collect the supernatant and add one milliliter of clean, wet amylose resin. Rotate the mixture for two hours at four degrees Celsius.Then load the protein amylose mixture in gravity flow chromatography columns. Wash the protein bound amylose resin with 10 times the bed volume of ice cold buffer C, followed by 10 bed volumes of ice cold buffer D.Elute eTRPV1 protein with 0.5 milliliter fractions up to five milliliters of ice cold degassed buffer D supplemented with 20 millimolar maltose. Perform the washes and elution at four degrees Celsius.Using a 100 kilodalton cutoff centrifugal filter unit, concentrate the eTRPV1 containing fractions to between two and 2.5 milligrams per milliliter for labeling. Next, add a ten-fold molar excess of MTSSL spin label from a 100 millimolar stock solution in DMSO to the concentrated protein. Repeat this twice every 30 minutes.Keep the reaction in the dark at room temperature for one hour and 30 minutes, followed by overnight incubation at four degrees Celsius. Then, load the spin label to eTRPV1 on a size-exclusion chromatography column equilibrated in buffer E, controlled by a fast protein liquid chromatography system. Collect the fractions containing eTRPV1 tetramer.Dry 10 milligrams of asolectin using a rotary evaporator under vacuum for one hour at 40 degrees Celsius. To make asolectin liposomes, add one milliliter of buffer F and sonicate the mixture for 15 minutes or until the sample is homogenous. Destabilize the liposomes by adding DDM to a final concentration of two millimolar and incubate for 30 minutes at room temperature.After adjusting the mixture to the DDM critical mycel concentration, as detailed in the text protocol, double the volume of the protein liposome mixture with buffer F.Remove the detergent by sequentially adding three aliquots of non-polar polystyrene absorbent beads at one hour intervals with gentle agitation at room temperature. Load the mixture onto a column filter to remove the beads and transfer the proteoliposome mixture to an ultra-centrifuge tube. Centrifuge the samples at 100, 000 times G for one hour at four degrees Celsius.Discard the supernatant and resuspend the pellet in 30 microliters of buffer F, and then perform spectroscopic measurements and analysis. The first checkpoint is the functional analysis of single-cysteine mutants before undertaking expression and purification protocols. Here, hek293 cells expressing wild type TRPV1, eTRPV1, and mutants were analyzed for capsaicin evoked responses using florescence calcium imaging.The second checkpoint is the biochemical characterization of spin labeled single-cysteine mutants through size-exclusion chromatography before proceeding with reconstitution and spectroscopic analysis. The third checkpoint is the functional characterization of reconstituted spin labeled single-cysteine mutants before undertaking spectroscopic analysis. Current voltage relationships determined by two electrode voltage clamp from Xenopus oocytes micro-injected with proteoliposomes containing spin labeled A702C were challenged with pH5 and blocked by capsazepine.Finally, it is possible to obtain the mobilities of spin labeled eTRPV1 mutants determined by continuous wave EPR. The first derivative spectra are shown for spin labeled cysteine mutants in DDM solution, and reconstituted in asolectin liposomes. Distances and mobilities of spin labeled eTRPV1 mutants in detergent were also determined by DEER.DEER echo and distance distribution of spin labeled mutant E651C is shown. After watching this video, you should have a good understanding of how to express, beautify, and reconstitute a mammalian ion channel for spectroscopic analysis. Generally, individuals new to this method will struggle because of the lack of protein function when removing cysteine residues, low protein yield, and protein instability either in purification and after spin labeling, and protein aggregation in detergent or liposomes.After its development, these protocols will allow obtaining distance and dynamic changes to explore mechanistic models for TRPV1 polymodal gating.

purification-and-reconstitution-of-trpv1-for-spectroscopic-analysis

Research

JoVE Journal

Biochemistry

Method for Efficient Refolding and Purification of Chemoreceptor Ligand Binding Domain

Identification of natural ligands of chemoreceptors and structural studies aimed at elucidation of the molecular basis of the ligand specificity can be greatly facilitated by the production of milligram amounts of pure, folded ligand binding domains. Attempts to heterologously express periplasmic ligand binding domains of bacterial chemoreceptors in Escherichia coli (E. coli) often result in their targeting into inclusion bodies. Here, a method is presented for protein recovery from inclusion bodies, its refolding and purification, using the periplasmic dCACHE ligand binding domain of Campylobacter jejuni (C. jejuni) chemoreceptor Tlp3 as an example. The approach involves expression of the protein of interest with a cleavable His6-tag, isolation and urea-mediated solubilisation of inclusion bodies, protein refolding by urea depletion, and purification by means of affinity chromatography, followed by tag removal and size-exclusion chromatography. The circular dichroism spectroscopy is used to confirm the folded state of the pure protein. It has been demonstrated that this protocol is generally useful for production of milligram amounts of dCACHE periplasmic ligand binding domains of other bacterial chemoreceptors in a soluble and crystallisable form.

A procedure is presented for the refolding of the dCACHE periplasmic ligand binding domain of Campylobacter jejuni chemoreceptor Tlp3 from inclusion bodies and the purification to yield milligram quantities of protein.

Identification of natural ligands of chemoreceptors and structural studies aimed at elucidation of the molecular basis of the ligand specificity can be ...

2 in 1: One-step Affinity Purification for the Parallel Analysis of Protein-Protein and Protein-Metabolite Complexes

Cellular processes are regulated by interactions between biological molecules such as proteins, metabolites, and nucleic acids. While the investigation of protein-protein interactions (PPI) is no novelty, experimental approaches aiming to characterize endogenous protein-metabolite interactions (PMI) constitute a rather recent development. Herein, we present a protocol that allows simultaneous characterization of the PPI and PMI of a protein of choice, referred to as bait. Our protocol was optimized for Arabidopsis cell cultures and combines affinity purification (AP) with mass spectrometry (MS)-based protein and metabolite detection. In short, transgenic Arabidopsis lines, expressing bait protein fused to an affinity tag, are first lysed to obtain a native cellular extract. Anti-tag antibodies are used to pull down protein and metabolite partners of the bait protein. The affinity-purified complexes are extracted using a one-step methyl tert-butyl ether (MTBE)/methanol/water method. Whilst metabolites separate into either the polar or the hydrophobic phase, proteins can be found in the pellet. Both metabolites and proteins are then analyzed by ultra-performance liquid chromatography-mass spectrometry (UPLC-MS or UPLC-MS/MS). Empty-vector (EV) control lines are used to exclude false positives. The major advantage of our protocol is that it enables identification of protein and metabolite partners of a target protein in parallel in near-physiological conditions (cellular lysate). The presented method is straightforward, fast, and can be easily adapted to biological systems other than plant cell cultures.

Protein-protein and protein-metabolite interactions are crucial for all cellular functions. Herein, we describe a protocol that allows parallel analysis of these interactions with a protein of choice. Our protocol was optimized for plant cell cultures and combines affinity purification with mass spectrometry-based protein and metabolite detection.

Cellular processes are regulated by interactions between biological molecules such as proteins, metabolites, and nucleic acids. While the investigation of ...

In Vitro Reconstitution of Self-Organizing Protein Patterns on Supported Lipid Bilayers

Many aspects of the fundamental spatiotemporal organization of cells are governed by reaction-diffusion type systems. In vitro reconstitution of such systems allows for detailed studies of their underlying mechanisms which would not be feasible in vivo. Here, we provide a protocol for the in vitro reconstitution of the MinCDE system of Escherichia coli, which positions the cell division septum in the cell middle. The assay is designed to supply only the components necessary for self-organization, namely a membrane, the two proteins MinD and MinE and energy in the form of ATP. We therefore fabricate an open reaction chamber on a coverslip, on which a supported lipid bilayer is formed. The open design of the chamber allows for optimal preparation of the lipid bilayer and controlled manipulation of the bulk content. The two proteins, MinD and MinE, as well as ATP, are then added into the bulk volume above the membrane. Imaging is possible by many optical microscopies, as the design supports confocal, wide-field and TIRF microscopy alike. In a variation of the protocol, the lipid bilayer is formed on a patterned support, on cell-shaped PDMS microstructures, instead of glass. Lowering the bulk solution to the rim of these compartments encloses the reaction in a smaller compartment and provides boundaries that allow mimicking of in vivo oscillatory behavior. Taken together, we describe protocols to reconstitute the MinCDE system both with and without spatial confinement, allowing researchers to precisely control all aspects influencing pattern formation, such as concentration ranges and addition of other factors or proteins, and to systematically increase system complexity in a relatively simple experimental setup.

In Vitro Reconstitution of Self-Organizing Protein Patterns on Supported Lipid Bilayers

We provide a protocol for in vitro self-organization assays of MinD and MinE on a supported lipid bilayer in an open chamber. Additionally, we describe how to enclose the assay in lipid-clad PDMS microcompartments to mimic in vivo conditions by reaction confinement.

Many aspects of the fundamental spatiotemporal organization of cells are governed by reaction-diffusion type systems. In vitro reconstitution of such ...

Anaerobic Protein Purification and Kinetic Analysis via Oxygen Electrode for Studying DesB Dioxygenase Activity and Inhibition

Oxygen-sensitive proteins, including those enzymes which utilize oxygen as a substrate, can have reduced stability when purified using traditional aerobic purification methods. This manuscript illustrates the technical details involved in the anaerobic purification process, including the preparation of buffers and reagents, the methods for column chromatography in a glove box, and the desalting of the protein prior to kinetics. Also described are the methods for preparing and using an oxygen electrode to perform kinetic characterization of an oxygen-utilizing enzyme. These methods are illustrated using the dioxygenase enzyme DesB, a gallate dioxygenase from the bacterium Sphingobium sp. strain SYK-6.

Here we present a protocol for anaerobic protein purification, anaerobic protein concentration, and subsequent kinetic characterization using an oxygen electrode system. The method is illustrated using the enzyme DesB, a dioxygenase enzyme which is more stable and active when purified and stored in an anaerobic environment.

Oxygen-sensitive proteins, including those enzymes which utilize oxygen as a substrate, can have reduced stability when purified using traditional aerobic ...

Expression, Purification, Crystallization, and Enzyme Assays of Fumarylacetoacetate Hydrolase Domain-Containing Proteins

Fumarylacetoacetate hydrolase (FAH) domain-containing proteins (FAHD) are identified members of the FAH superfamily in eukaryotes. Enzymes of this superfamily generally display multi-functionality, involving mainly hydrolase and decarboxylase mechanisms. This article presents a series of consecutive methods for the expression and purification of FAHD proteins, mainly FAHD protein 1 (FAHD1) orthologues among species (human, mouse, nematodes, plants, etc.). Covered methods are protein expression in E. coli, affinity chromatography, ion exchange chromatography, preparative and analytical gel filtration, crystallization, X-ray diffraction, and photometric assays. Concentrated protein of high levels of purity (&#62;98%) may be employed for crystallization or antibody production. Proteins of similar or lower quality may be employed in enzyme assays or used as antigens in detection systems (Western-Blot, ELISA). In the discussion of this work, the identified enzymatic mechanisms of FAHD1 are outlined to describe its hydrolase and decarboxylase bi-functionality in more detail.

Expression and purification of fumarylacetoacetate hydrolase domain-containing proteins is described with examples (expression in E. coli, FPLC). Purified proteins are used for crystallization and antibody production and employed for enzyme assays. Selected photometric assays are presented to display the multi-functionality of FAHD1 as oxaloacetate decarboxylase and acylpyruvate hydrolase.

Fumarylacetoacetate hydrolase (FAH) domain-containing proteins (FAHD) are identified members of the FAH superfamily in eukaryotes. Enzymes of this ...

Detergent-assisted Reconstitution of Recombinant Drosophila Atlastin into Liposomes for Lipid-mixing Assays

Membrane fusion is a crucial process in the eukaryotic cell. Specialized proteins are necessary to catalyze fusion. Atlastins are endoplasmic reticulum (ER) resident proteins implicated in homotypic fusion of the ER. We detail here a method for purifying a glutathione S-transferase (GST) and poly-histidine tagged Drosophila atlastin by two rounds of affinity chromatography. Studying fusion reactions in vitro requires purified fusion proteins to be inserted into a lipid bilayer. Liposomes are ideal model membranes, as lipid composition and size may be adjusted. To this end, we describe a reconstitution method by detergent removal for Drosophila atlastin into preformed liposomes. While several reconstitution methods are available, reconstitution by detergent removal has several advantages that make it suitable for atlastins and other similar proteins. The advantage of this method includes a high reconstitution yield and correct orientation of the reconstituted protein. This method can be extended to other membrane proteins and for other applications that require proteoliposomes. Additionally, we describe a FRET based lipid mixing assay of proteoliposomes used as a measurement of membrane fusion.

Detergent-assisted Reconstitution of Recombinant Drosophila Atlastin into Liposomes for Lipid-mixing Assays

Biological membrane fusion is catalyzed by specialized fusion proteins. Measuring the fusogenic properties of proteins can be achieved by lipid mixing assays. We present a method for purifying recombinant Drosophila atlastin, a protein that mediates homotypic fusion of the ER, reconstituting it to preformed liposomes, and testing for fusion capacity.

Membrane fusion is a crucial process in the eukaryotic cell. Specialized proteins are necessary to catalyze fusion. Atlastins are endoplasmic reticulum ...

Isolation of Active Caenorhabditis elegans Nuclear Extract and Reconstitution for In Vitro Transcription

Caenorhabditis elegans has been an important model system for biological research since it was introduced in 1963. However, C. elegans has not been fully utilized in the biochemical study of biological reactions using its nuclear extracts such as in vitro transcription and DNA replication. A significant hurdle for using C. elegans in biochemical studies is disrupting the nematode&#39;s thick outer cuticle without sacrificing the activity of the nuclear extract. While several methods are used to break the cuticle, such as Dounce homogenization or sonication, they often lead to protein instability. There are no established protocols for isolating active nuclear proteins from larva or adult C. elegans for in vitro reactions. Here, the protocol describes in detail the homogenization of larval stage 4 C. elegans using a Balch homogenizer. The Balch homogenizer uses pressure to slowly force the animals through a narrow gap breaking the cuticle in the process. The uniform design and precise machining of the Balch homogenizer allow for consistent grinding of animals between experiments. Fractionating the homogenate obtained from the Balch homogenizer yields functionally active nuclear extract that can be used in an in vitro method for assaying transcription activity of C. elegans.

Isolation of Active Caenorhabditis elegans Nuclear Extract and Reconstitution for In Vitro Transcription

Here, we describe a detailed protocol for isolating active nuclear extract from larval stage 4 C. elegans and visualizing transcription activity in an in vitro system.

Caenorhabditis elegans has been an important model system for biological research since it was introduced in 1963. However, C. elegans has not been fully ...

Reconstitution of Msp1 Extraction Activity with Fully Purified Components

As the center for oxidative phosphorylation and apoptotic regulation, mitochondria play a vital role in human health. Proper mitochondrial function depends on a robust quality control system to maintain protein homeostasis (proteostasis). Declines in mitochondrial proteostasis have been linked to cancer, aging, neurodegeneration, and many other diseases. Msp1 is a AAA+ ATPase anchored in the outer mitochondrial membrane that maintains proteostasis by removing mislocalized tail-anchored proteins. Using purified components reconstituted into proteoliposomes, we have shown that Msp1 is necessary and sufficient to extract a model tail-anchored protein from a lipid bilayer. Our simplified reconstituted system overcomes several of the technical barriers that have hindered detailed study of membrane protein extraction. Here, we provide detailed methods for the generation of liposomes, membrane protein reconstitution, and the Msp1 extraction assay.

Here, we present a detailed protocol for reconstitution of Msp1 extraction activity with fully purified components in defined proteoliposomes.

As the center for oxidative phosphorylation and apoptotic regulation, mitochondria play a vital role in human health. Proper mitochondrial function ...

Purification of Endogenous Drosophila Transient Receptor Potential Channels

Drosophila phototransduction is one of the fastest known G protein-coupled signaling pathways. To ensure the specificity and efficiency of this cascade, the calcium (Ca2+)-permeable cation channel, transient receptor potential (TRP), binds tightly to the scaffold protein, inactivation-no-after-potential D (INAD), and forms a large signaling protein complex with eye-specific protein kinase C (ePKC) and phospholipase C&#946;/No receptor potential A (PLC&#946;/NORPA). However, the biochemical properties of the Drosophila TRP channel remain unclear. Based on the assembling mechanism of INAD protein complex, a modified affinity purification plus competition strategy was developed to purify the endogenous TRP channel. First, the purified histidine (His)-tagged NORPA 863-1095 fragment was bound to Ni-beads and used as bait to pull down the endogenous INAD protein complex from Drosophila head homogenates. Then, excessive purified glutathione S-transferase (GST)-tagged TRP 1261-1275 fragment was added to the Ni-beads to compete with the TRP channel. Finally, the TRP channel in the supernatant was separated from the excessive TRP 1261-1275 peptide by size-exclusion chromatography. This method makes it possible to study the gating mechanism of the Drosophila TRP channel from both biochemical and structural angles. The electrophysiology properties of purified Drosophila TRP channels can also be measured in the future.

Purification of Endogenous Drosophila Transient Receptor Potential Channels

Based on the assembling mechanism of the INAD protein complex, in this protocol, a modified affinity purification plus competition strategy was developed to purify the endogenous Drosophila TRP channel.

Drosophila phototransduction is one of the fastest known G protein-coupled signaling pathways. To ensure the specificity and efficiency of this cascade, ...

Extraction and Purification of FAHD1 Protein from Swine Kidney and Mouse Liver

Fumarylacetoacetate hydrolase domain-containing protein 1 (FAHD1) is the first identified member of the FAH superfamily in eukaryotes, acting as oxaloacetate decarboxylase in mitochondria. This article presents a series of methods for the extraction and purification of FAHD1 from swine kidney and mouse liver. Covered methods are ionic exchange chromatography with fast protein liquid chromatography (FPLC), preparative and analytical gel filtration with FPLC, and proteomic approaches. After total protein extraction, ammonium sulfate precipitation and ionic exchange chromatography were explored, and FAHD1 was extracted via a sequential strategy using ionic exchange and size-exclusion chromatography. This representative approach may be adapted to other proteins of interest (expressed at significant levels) and modified for other tissues. Purified protein from tissue may support the development of high-quality antibodies, and/or potent and specific pharmacological inhibitors.

This protocol describes how to extract fumarylacetoacetate hydrolase domain-containing protein 1 (FAHD1) from swine kidney and mouse liver. The listed methods may be adapted to other proteins of interest and modified for other tissues.