This work was supported by grant no. 2016/23/D/NZ3/01314 from the National Science Centre (NCN), Krakow, Poland.

1. Generation of expression plasmids
<ol>
	<li>Examine membrane topology of the proteins of interest using a topology predicting tool.</li>
	<li>Design the cloning strategy so that the larger and smaller fragments would face the cytoplasm once the fusion proteins have been inserted into Golgi membranes. If, as in the case&#160;presented here, both N- and C-termini of the proteins of interest are cytoplasmically oriented, tag the proteins in eight possible ways (see Figure 1B). If N- or C-terminus of one or both proteins of interest is luminally oriented, exclude it from tagging.</li>
	<li>Subclone the genes of interest into appropriate expression vectors (see Table of Materials) by following standard cloning protocols. 
	NOTE: Testing all the possible orientations is recommended, since some tagging options may not work due to insufficient proximity, suboptimal orientation, or spatial constraints.</li>
</ol>
2. Transient transfection of the expression plasmids into the cells
<ol>
	<li>Harvest the adherent HEK293T cell culture by trypsinization and resuspend the cells in a dedicated complete growth medium. Plate the cells (2 x 104/100 &#181;L/well) onto a clear bottom, white side 96-well plate. Adjust the total number of wells to accommodate all tested combinations and controls including replicates. 
	NOTE: Attempt to use only the inner 60 wells of the plate to minimize thermal shifts and avoid overnight evaporation. Using poly-D-lysine-coated plates is highly recommended such as the ones indicated in the Table of Materials, otherwise cells may detach during the subsequent washing steps.</li>
	<li>Culture the cells overnight in standard conditions (37 &#176;C, 5% CO2).</li>
	<li>On the next day transfect the cells with the desired combinations of expression plasmids obtained in point 1.1.
	<ol>
		<li>Dilute expression plasmids in a serum-free medium (see Table of Materials) to 6.25 ng/&#181;L for each construct.</li>
		<li>Add the lipid-based transfection reagent at an appropriate lipid-to-DNA ratio and incubate according to the manufacturer&#8217;s instruction.</li>
		<li>Add 8 &#181;L of lipid:DNA mixture to designated wells. Mix the content of the plate by gentle rotation. This results in transfection of both expression constructs at 50 ng/well.</li>
	</ol>
	</li>
	<li>Culture the cells for 20-24 h in standard conditions (37 &#176;C, 5% CO2). 
	NOTE: Culturing the cells for a longer time may result in higher levels of fusion protein expression, which may promote a non-specific association between the fragments.</li>
</ol>
3. Medium exchange
<ol>
	<li>On the next day replace the conditioned medium with 100 &#181;L of a serum-free medium in each well. Make sure that the cells have not detached upon medium exchange. 
	NOTE: This step should be done 2-3 h before the addition of the furimazine working solution. Serum withdrawal minimizes background caused by autoluminescence of furimazine.</li>
</ol>
4. Preparation of furimazine working solution
<ol>
	<li>Just before the measurement, mix 1 volume of furimazine with 19 volumes of a dilution buffer (a 20-fold dilution). 
	NOTE: The total volume of the furimazine working solution to be prepared depends on the number of individual wells to be analyzed (the furimazine working solution is added to the cell culture medium in a 1:5 ratio, therefore, to each well previously filled with 100 &#181;L of a serum-free medium 25 &#181;L of the furimazine working solution should be added).</li>
	<li>Add the furimazine working solution to designated wells (25 &#181;L/well). Gently mix the plate by hand or using an orbital shaker (e.g., 15 s at 300-500 rpm).</li>
</ol>
5. Measuring luminescence
<ol>
	<li>Insert the plate into a luminescence microplate reader.
	<ol>
		<li>For experiments that are to be performed at 37 &#176;C equilibrate the plate for 10-15 min at the indicated temperature.</li>
	</ol>
	</li>
	<li>Select the wells to be analyzed.</li>
	<li>Read luminescence with integration time of 0.3 s. Continue to monitor luminescence for up to 2 h when required.</li>
</ol>
6. Data analysis
<ol>
	<li>Calculate mean values and standard deviations for all the tested and control combinations.</li>
	<li>Analyze data using one-way ANOVA with multiple comparisons.</li>
	<li>Calculate fold change values by dividing a mean luminescence obtained for combinations of interest by a mean luminescence obtained for the corresponding negative controls. Evaluate the results. 
	NOTE: The approach to data analysis proposed here assumes that the interaction can be claimed if the luminescence obtained for the tested combination is statistically significantly higher than the luminescence obtained for the corresponding control combination and, at the same time, the ratio of these two values exceeds 10.</li>
</ol>

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B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+and+purification+of+the+rat+liver+Golgi+membrane+UDP-N-acetylgalactosamine+transporter.">Identification and purification of the rat liver Golgi membrane UDP-N-acetylgalactosamine transporter.</a> Journal of Biological Chemistry. 274 (7), 4474-4479 (1999).</li><li>Gao, X., Dean, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distinct+protein+domains+of+the+yeast+Golgi+GDP-mannose+transporter+mediate+oligomer+assembly+and+export+from+the+endoplasmic+reticulum.">Distinct protein domains of the yeast Golgi GDP-mannose transporter mediate oligomer assembly and export from the endoplasmic reticulum.</a> Journal of Biological Chemistry. 275 (23), 17718-17727 (2000).</li><li>Olczak, M., Guillen, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+a+mutation+and+an+alternative+splicing+of+UDP-galactose+transporter+in+MDCK-RCAr+cell+line.">Characterization of a mutation and an alternative splicing of UDP-galactose transporter in MDCK-RCAr cell line.</a> Biochimica et Biophysica Acta. 1763 (1), 82-92 (2006).</li><li>Maszczak-Seneczko, D., Sosicka, P., Majkowski, M., Olczak, T., Olczak, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=UDP-N-acetylglucosamine+transporter+and+UDP-galactose+transporter+form+heterologous+complexes+in+the+Golgi+membrane.">UDP-N-acetylglucosamine transporter and UDP-galactose transporter form heterologous complexes in the Golgi membrane.</a> FEBS Letters. 586 (23), 4082-4087 (2012).</li><li>Nji, E., Gulati, A., Qureshi, A. 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The authors have nothing to disclose.

Here we provide a detailed protocol enabling the demonstration of heterologous complexes formed between Golgi-resident type III membrane proteins, such as NSTs, using the split luciferase complementation assay. The proposed approach to data analysis and interpretation involves relating the luminescence obtained for the protein combination of interest to the luminescence obtained for the corresponding control combination, which is composed of one of the proteins of interest fused with the larger fragment and the control p...

This manuscript describes a step-by-step protocol to check for the presence of heterologous interactions between Golgi-resident type III membrane proteins in transiently transfected human cells using the most recent variant of the split luciferase complementation assay. The procedure has been most extensively tested against nucleotide sugar transporters (NSTs) but we were also able to obtain positive results for other Golgi-resident type III membrane proteins whose N- and/or C-termini are facing the cytoplasm.
Our research group explores the role of NSTs in glycosylation of macromolecules. NSTs are Golgi- and/or ER-resident type III membrane proteins with N- and C-termini facing the cytoplasmic side of the organellar membrane1. NSTs are thought to carry nucleotide-activated sugars across organelle membranes to supply glycosyltransferases with their substrates. NSTs form dimers&#160;and/or higher oligomers2,3,4,5,6,7,8,9,10. Moreover, heterologous interactions between&#160;different NSTs have also been reported6,11. NSTs were also demonstrated to form complexes with functionally related glycosylation enzymes12,13,14. We sought for an alternative to the presently used technique, fluorescence lifetime imaging (FLIM)-based FRET approach, for studying interactions of NSTs and functionally related Golgi-resident proteins, so we decided to test the split&#160;luciferase complementation assay. It allowed us to identify a novel interaction between an NST and a functionally related glycosylation enzyme9.
The most recent modification of the split luciferase complementation assay, NanoBiT, is used in the protocol presented here15. It relies on the reconstitution of the luciferase enzyme (e.g., NanoLuc) from the two fragments - the large one, termed as large BiT or LgBiT, a 17.6 kDa protein, and the small one, composed of only 11 amino acids, termed as small BiT or SmBiT. The two proteins of interest are fused with the complementary fragments and transiently expressed in a human cell line. If the two fusion proteins interact, a luminescence is produced in situ upon addition of a cell-permeable substrate. These two fragments have been optimized so that they associate with minimum affinity unless being brought together by an interaction between the proteins of interest they are fused to.
In general, bioluminescence-based methods have some advantages over the ones based on fluorescence. Bioluminescent signals have a higher signal-to-noise ratio because the background luminescence is negligible compared to the luciferase-derived signal16. In contrast, fluorescence-based approaches usually suffer from a relatively high background caused by the phenomenon of autofluorescence. Besides, bioluminescence is less detrimental to the analyzed cells than fluorescence, as in the former case there is no need to excite the sample. For those reasons bioluminescent approaches to studying PPIs in vivo outcompete the commonly used fluorescent methods like F&#246;rster Resonance Energy Transfer (FRET) or Bimolecular Fluorescence Complementation (BiFC).
Our protocol relies on referring luminescence obtained for the protein combination of interest to luminescence obtained for the control combination. The latter includes the one of the tested proteins which is fused with a larger fragment and a control&#160;protein (e.g., HaloTag),&#160;fused with a smaller fragment. The latter&#160;is a protein of bacterial origin that is not expected to interact with any of mammalian proteins. Using this protein as a control poses limitations to the topology of the Golgi-resident pairs of proteins to be analyzed. Since in mammalian cells this protein is synthesized in the cytoplasm, both proteins of interest should have at least one cytoplasmic tail.
This approach can be particularly useful for initial screening of PPIs. It may become the method of choice when the fusion proteins of interest are expressed at levels that are simply insufficient for other approaches to be applied. Similarly, the split luciferase complementation assay can be the best option if the proteins of interest are expressed at high levels, but this adversely affects their subcellular localization or is known to force non-specific interactions. Since the smaller fragment has only 11 amino acids, the split luciferase complementation assay can be applied when using larger tags is impossible. Finally, it can be employed to further confirm data obtained using other techniques, as in the case&#160;presented here.

<table>
	<tbody>
		<tr>
			<td>0.25% trypsin-EDTA solution</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Adherent mammalian cell line</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>BioCoat Poly-D-Lysine 96-well White/Clear Flat Bottom TC-treated Microplate, with Lid</td>
			<td>Corning</td>
			<td>356651</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell culture centrifuge</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cell culture supplements (heat-inactivated fetal bovine serum, L-glutamine, penicillin, streptamycin)</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>CO2 incubator</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Expression plasmids encoding protein(s) of interest not tagged with NanoBiT fragments</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>FuGENE HD Transfection Reagent</td>
			<td>Promega</td>
			<td>E2311</td>
			<td></td>
		</tr>
		<tr>
			<td>GloMax Discover Microplate Reader (or a different luminescence microplate reader)</td>
			<td>Promega</td>
			<td>GM3000</td>
			<td></td>
		</tr>
		<tr>
			<td>Growth medium dedicated to the cell line used</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Materials and reagents for standard molecular cloning (bacteria, thermostable polymerase, restriction enzymes, DNA ligase, materials and reagents for nucleic acid purification)</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>NanoBiT MCS Starter System</td>
			<td>Promega</td>
			<td>N2014</td>
			<td>This kit contains vectors enabling tagging of the proteins of interest with NanoBiT fragments at different orientations as well as the control plasmid encoding HaloTag protein fused with SmBiT and a positive control plasmid pair.</td>
		</tr>
		<tr>
			<td>Nano-Glo Live Cell Assay System</td>
			<td>Promega</td>
			<td>N2011</td>
			<td>This kit contains furimazine, which is a substrate enabling detection of the NanoLuc activity in living cells, and a dedicated dilution buffer.</td>
		</tr>
		<tr>
			<td>Opti-MEM I Reduced Serum Medium, no phenol red</td>
			<td>Gibco</td>
			<td>11058021</td>
			<td></td>
		</tr>
		<tr>
			<td>Oribital shaker</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Software for data analysis (e.g. GraphPad Prism)</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Thermocycler</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

demonstration of heterologous complexes formed by golgi-resident type iii membrane proteins using split luciferase complementation assay

The goal of this protocol is to explore the applicability of the most recent variant of split luciferase complementation for demonstrating heterologous complexes formed by nucleotide sugar transporters (NSTs). These ER- and Golgi-resident multitransmembrane proteins carry the cytoplasmically synthesized nucleotide sugars across organelle membranes to supply enzymes that mediate glycosylation with their substrates. NSTs exist as dimers and/or higher oligomers. Heterologous interactions between&#160;different NSTs have also been reported. To verify whether the technique is suitable for studying the phenomenon of NST heteromerization, we tested it against a combination of the two Golgi-resident NSTs that have been previously shown to associate by several other means. The&#160;luciferase complementation assay appears to be particularly suitable for studying interactions between Golgi-resident membrane proteins, as it does not require high expression levels, which often trigger protein mislocalization and increase the risk of false positives.

To obtain the most reliable data in this approach all the possible combinations should be tested (see Figure 1). In parallel, positive and negative controls should be included. The positive control should consist of the two proteins that are known to interact, of which one is fused with the larger fragment and the other is fused with the smaller fragment. The negative control ideally should consist of the two non-interacting type III membrane proteins tagged likewise. However, establishing s...

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Demonstration of Heterologous Complexes formed by Golgi-Resident Type III Membrane Proteins using Split Luciferase Complementation Assay

The&#160;protocol presented here is intended to demonstrate the occurrence of heterologous interactions between Golgi-resident type III membrane proteins with cytoplasmically exposed N- and/or C-termini in live mammalian cells using the most recent variant of the split luciferase complementation assay.

The goal of this protocol is to explore the applicability of the most recent variant of split luciferase complementation for demonstrating heterologous ...

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2.2K Views.  University of Wroclaw. The protocol presented here is intended to demonstrate the occurrence of heterologous interactions between Golgi-resident type III membrane proteins with cytoplasmically exposed N- and/or C-termini in live mammalian cells using the most recent variant of the split luciferase complementation assay.

Video: Demonstration of Heterologous Complexes formed by Golgi-Resident Type III Membrane Proteins using Split Luciferase Complementation Assay

Studying Membrane Biogenesis with a Luciferase-Based Reporter Gene Assay

To study the coordination of different lipid synthesis pathways during membrane biogenesis it is useful to work with an experimental system where membrane biogenesis occurs rapidly and in an inducible manner. We have found that phagocytosis of latex beads is practical for these purposes as cells rapidly synthesize membrane lipids to replenish membrane pools lost as  wrapping material  during particle engulfment. Here, we describe procedures for studying changes in phagocytosis-induced gene expression with a luciferase-based reporter gene approach using the Dual-Glo  Luciferase Assay System from Promega.

Here, we describe procedures for studying changes in phagocytosis-induced gene expression with a luciferase-based reporter gene approach using the Dual-GloTM Luciferase Assay System from Promega.

To study the coordination of different lipid synthesis pathways during membrane biogenesis it is useful to work with an experimental system where membrane ...

Antibody Profiling by Luciferase Immunoprecipitation Systems (LIPS)

Technologies for comprehensively understanding and quantifying antibody profiles to autoantigens and infectious agents may yield new insights into disease mechanisms and may elucidate new markers to substratify disease with different clinical features and better understand pathogenesis. We have developed a highly quantitative method called Luciferase Immunoprecipitation Systems (LIPS) for profiling patient sera antibody responses to autoantigens and pathogen antigens associated with infection. Unlike ELISAs, the highly sensitive LIPS is easily implemented to survey humoral serological response profiles to different antigens in a universal format and produces dynamic antibody titer ranges up to 6 log10 for some antigens. In these studies, quantitative profiling by LIPS of patient humoral responses against panels of antigens or even the entire proteome of some pathogens (i.e. HIV), is typically more informative than testing a single antigen by ELISA. In addition, LIPS also eliminates time and effort needed to produce highly purified antigens as well as the labor-intensive assay optimization steps needed for standard ELISAs. Here we provide a detailed protocol describing the technical aspects of performing LIPS assays for readily profiling antibody responses to single or multiple antigens.

Antibody Profiling by Luciferase Immunoprecipitation Systems LIPS

The technical aspects of performing LIPS (Luciferase Immunoprecipitation Systems) are described. The overall approach involves expressing chimeric genes encoding antigens fused to Renilla luciferase (Ruc) in mammalian cells. Crude Ruc-antigen extracts are then prepared and, without purification, employed in immunoprecipitation assays to quantify antibodies.

Technologies for comprehensively understanding and quantifying antibody profiles to autoantigens and infectious agents may yield new insights into disease ...

Crystallizing Membrane Proteins for Structure Determination using Lipidic Mesophases

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the lipidic cubic phase or in meso method. The method has been shown to be quite versatile in that it has been used to solve X-ray crystallographic structures of prokaryotic and eukaryotic proteins, proteins that are monomeric, homo- and hetero-multimeric, chromophore-containing and chromophore-free, and alpha-helical and beta-barrel proteins. Recent successes using in meso crystallization are the human engineered beta2-adrenergic and adenosine A2a G protein-coupled receptors. Protocols are presented for reconstituting the membrane protein into the monoolein-based mesophase, and for setting up crystallizations in the manual mode. Additional steps in the overall process, such as crystal harvesting, are to be addressed in future video articles. The time required to prepare the protein-loaded mesophase and to set up a crystallization plate manually is about one hour.

Herein is described the procedure implemented in the Caffrey Membrane Structural and Functional Biology Group to set up manually crystallization trials of membrane proteins in lipidic mesophases.

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the ...

Identification of Growth Inhibition Phenotypes Induced by Expression of Bacterial Type III Effectors in Yeast

Many Gram-negative pathogenic bacteria use a type III secretion system to translocate a suite of effector proteins into the cytosol of host cells. Within the cell, type III effectors subvert host cellular processes to suppress immune responses and promote pathogen growth. Numerous type III effectors of plant and animal bacterial pathogens have been identified to date, yet only a few of them are well characterized. Understanding the functions of these effectors has been undermined by a combination of functional redundancy in the effector repertoire of a given bacterial strain, the subtle effects that they may exert to increase virulence, roles that are possibly specific to certain infection stages, and difficulties in genetically manipulating certain pathogens. Expression of type III effectors in the budding yeast Saccharomyces cerevisiae may allow circumventing these limitations and aid to the functional characterization of effector proteins. Because type III effectors often target cellular processes that are conserved between yeast and other eukaryotes, their expression in yeast may result in growth inhibition phenotypes that can be exploited to elucidate effector functions and targets. Additional advantages to using yeast for functional studies of bacterial effectors include their genetic tractability, information on predicted functions of the vast majority of their ORFs, and availability of numerous tools and resources for both genome-wide and small-scale experiments. Here we discuss critical factors for designing a yeast system for the expression of bacterial type III effector proteins. These include an appropriate promoter for driving expression of the effector gene(s) of interest, the copy number of the effector gene, the epitope tag used to verify protein expression, and the yeast strain. We present procedures to induce expression of effectors in yeast and to verify their expression by immunoblotting. In addition, we describe a spotting assay on agar plates for the identification of effector-induced growth inhibition phenotypes. The use of this protocol may be extended to the study of pathogenicity factors delivered into the host cell by any pathogen and translocation mechanism.

In this video, we describe a procedure for the expression of bacterial type III effectors in yeast and the identification of effector-induced growth inhibition phenotypes. Such phenotypes can be subsequently exploited to elucidate effector functions and targets.

Many Gram-negative pathogenic bacteria use a type III secretion system to translocate a suite of effector proteins into the cytosol of host cells. Within ...

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 ...

High-throughput Crystallization of Membrane Proteins Using the Lipidic Bicelle Method

Membrane proteins (MPs) play a critical role in many physiological processes such as pumping specific molecules across the otherwise impermeable membrane bilayer that surrounds all cells and organelles. Alterations in the function of MPs result in many human diseases and disorders; thus, an intricate understanding of their structures remains a critical objective for biological research. However, structure determination of MPs remains a significant challenge often stemming from their hydrophobicity. 
MPs have substantial hydrophobic regions embedded within the bilayer. Detergents are frequently used to solubilize these proteins from the bilayer generating a protein-detergent micelle that can then be manipulated in a similar manner as soluble proteins. Traditionally, crystallization trials proceed using a protein-detergent mixture, but they often resist crystallization or produce crystals of poor quality. These problems arise due to the detergent&prime;s inability to adequately mimic the bilayer resulting in poor stability and heterogeneity. In addition, the detergent shields the hydrophobic surface of the MP reducing the surface area available for crystal contacts. To circumvent these drawbacks MPs can be crystallized in lipidic media, which more closely simulates their endogenous environment, and has recently become a de novo technique for MP crystallization.
Lipidic cubic phase (LCP) is a three-dimensional lipid bilayer penetrated by an interconnected system of aqueous channels1. Although monoolein is the lipid of choice, related lipids such as monopalmitolein and monovaccenin have also been used to make LCP2. MPs are incorporated into the LCP where they diffuse in three dimensions and feed crystal nuclei. A great advantage of the LCP is that the protein remains in a more native environment, but the method has a number of technical disadvantages including high viscosity (requiring specialized apparatuses) and difficulties in crystal visualization and manipulation3,4. Because of these technical difficulties, we utilized another lipidic medium for crystallization-bicelles5,6 (Figure 1). Bicelles are lipid/amphiphile mixtures formed by blending a phosphatidylcholine lipid (DMPC) with an amphiphile (CHAPSO) or a short-chain lipid (DHPC). Within each bicelle disc, the lipid molecules generate a bilayer while the amphiphile molecules line the apolar edges providing beneficial properties of both bilayers and detergents. Importantly, below their transition temperature, protein-bicelle mixtures have a reduced viscosity and are manipulated in a similar manner as detergent-solubilized MPs, making bicelles compatible with crystallization robots. 
Bicelles have been successfully used to crystallize several membrane proteins5,7-11 (Table 1). This growing collection of proteins demonstrates the versatility of bicelles for crystallizing both alpha helical and beta sheet MPs from prokaryotic and eukaryotic sources. Because of these successes and the simplicity of high-throughput implementation, bicelles should be part of every membrane protein crystallographer&prime;s arsenal. In this video, we describe the bicelle methodology and provide a step-by-step protocol for setting up high-throughput crystallization trials of purified MPs using standard robotics.

Bicelles are lipid/amphiphile mixtures that maintain membrane proteins (MPs) within a lipid bilayer but have unique phase behavior that facilitates high-throughput screening by crystallization robots. This technique has successfully produced a number of high-resolution structures from both prokaryotic and eukaryotic sources. This video describes protocols for generating the lipidic bicelle mixture, incorporating MPs into the bicelle mixture, setting up crystallizations trials (manually as well as robotically) and harvesting crystals from the medium.

Membrane proteins (MPs) play a critical role in many physiological processes such as pumping specific molecules across the otherwise impermeable membrane ...

Use of a Robot for High-throughput Crystallization of Membrane Proteins in Lipidic Mesophases

Structure-function studies of membrane proteins greatly benefit from having available high-resolution 3-D structures of the type provided through macromolecular X-ray crystallography (MX). An essential ingredient of MX is a steady supply of ideally diffraction-quality crystals. The in meso or lipidic cubic phase (LCP) method for crystallizing membrane proteins is one of several methods available for crystallizing membrane proteins. It makes use of a bicontinuous mesophase in which to grow crystals. As a method, it has had some spectacular successes of late and has attracted much attention with many research groups now interested in using it. One of the challenges associated with the method is that the hosting mesophase is extremely viscous and sticky, reminiscent of a thick toothpaste. Thus, dispensing it manually in a reproducible manner in small volumes into crystallization wells requires skill, patience and a steady hand. A protocol for doing just that was developed in the Membrane Structural &amp; Functional Biology (MS&amp;FB) Group1-3. JoVE video articles describing the method are available1,4. 
The manual approach for setting up in meso trials has distinct advantages with specialty applications, such as crystal optimization and derivatization. It does however suffer from being a low throughput method. Here, we demonstrate a protocol for performing in meso crystallization trials robotically. A robot offers the advantages of speed, accuracy, precision, miniaturization and being able to work continuously for extended periods under what could be regarded as hostile conditions such as in the dark, in a reducing atmosphere or at low or high temperatures. An in meso robot, when used properly, can greatly improve the productivity of membrane protein structure and function research by facilitating crystallization which is one of the slow steps in the overall structure determination pipeline. 
In this video article, we demonstrate the use of three commercially available robots that can dispense the viscous and sticky mesophase integral to in meso crystallogenesis. The first robot was developed in the MS&amp;FB Group5,6. The other two have recently become available and are included here for completeness.	
An overview of the protocol covered in this article is presented in Figure 1. All manipulations were performed at room temperature (~20 &deg;C) under ambient conditions.

Herein is described a robotic approach to high-throughput crystallization of membrane proteins in lipidic mesophases for use in structure determination using macromolecular X-ray crystallography. Three robots capable of handling the viscous and sticky protein-laden mesophase integral to the method are introduced.

Structure-function studies of membrane proteins greatly benefit from having available high-resolution 3-D structures of the type provided through ...

Harvesting and Cryo-cooling Crystals of Membrane Proteins Grown in Lipidic Mesophases for Structure Determination by Macromolecular Crystallography

An important route to understanding how proteins function at a mechanistic level is to have the structure of the target protein available, ideally at atomic resolution. Presently, there is only one way to capture such information as applied to integral membrane proteins (Figure 1), and the complexes they form, and that method is macromolecular X-ray crystallography (MX). To do MX diffraction quality crystals are needed which, in the case of membrane proteins, do not form readily. A method for crystallizing membrane proteins that involves the use of lipidic mesophases, specifically the cubic and sponge phases1-5, has gained considerable attention of late due to the successes it has had in the G protein-coupled receptor field6-21 (<a href="http://www.mpdb.tcd.ie" target="_blank">www.mpdb.tcd.ie</a>). However, the method, henceforth referred to as the in meso or lipidic cubic phase method, comes with its own technical challenges. These arise, in part, due to the generally viscous and sticky nature of the lipidic mesophase in which the crystals, which are often micro-crystals, grow. Manipulating crystals becomes difficult as a result and particularly so during harvesting22,23. Problems arise too at the step that precedes harvesting which requires that the glass sandwich plates in which the crystals grow (Figure 2)24,25 are opened to expose the mesophase bolus, and the crystals therein, for harvesting, cryo-cooling and eventual X-ray diffraction data collection.
The cubic and sponge mesophase variants (Figure 3) from which crystals must be harvested have profoundly different rheologies4,26. The cubic phase is viscous and sticky akin to a thick toothpaste. By contrast, the sponge phase is more fluid with a distinct tendency to flow. Accordingly, different approaches for opening crystallization wells containing crystals growing in the cubic and the sponge phase are called for as indeed different methods are required for harvesting crystals from the two mesophase types. Protocols for doing just that have been refined and implemented in the Membrane Structural and Functional Biology (MS&amp;FB) Group, and are described in detail in this JoVE article (Figure 4). Examples are given of situations where crystals are successfully harvested and cryo-cooled. We also provide examples of cases where problems arise that lead to the irretrievable loss of crystals and describe how these problems can be avoided. In this article the Viewer is provided with step-by-step instructions for opening glass sandwich crystallization wells, for harvesting and for cryo-cooling crystals of membrane proteins growing in cubic and in sponge phases.

Herein is described procedures implemented in the Caffrey Membrane Structural and Functional Biology Group to harvest and cryo-cool membrane protein crystals grown in lipidic cubic and sponge phases for use in structure determination using macromolecular X-ray crystallography.

An important route to understanding how proteins function at a mechanistic level is to have the structure of the target protein available, ideally at ...

High-throughput Functional Screening using a Homemade Dual-glow Luciferase Assay

We present a rapid and inexpensive high-throughput screening protocol to identify transcriptional regulators of alpha-synuclein, a gene associated with Parkinson&#39;s disease. 293T cells are transiently transfected with plasmids from an arrayed ORF expression library, together with luciferase reporter plasmids, in a one-gene-per-well microplate format. Firefly luciferase activity is assayed after 48 hr to determine the effects of each library gene upon alpha-synuclein transcription, normalized to expression from an internal control construct (a hCMV promoter directing Renilla luciferase). This protocol is facilitated by a bench-top robot enclosed in a biosafety cabinet, which performs aseptic liquid handling in 96-well format. Our automated transfection protocol is readily adaptable to high-throughput lentiviral library production or other functional screening protocols requiring triple-transfections of large numbers of unique library plasmids in conjunction with a common set of helper plasmids. We also present an inexpensive and validated alternative to commercially-available, dual luciferase reagents which employs PTC124, EDTA, and pyrophosphate to suppress firefly luciferase activity prior to measurement of Renilla luciferase. Using these methods, we screened 7,670 human genes and identified 68 regulators of alpha-synuclein. This protocol is easily modifiable to target other genes of interest.

We present a rapid and inexpensive screening method for identifying transcriptional regulators using high-throughput robotic transfections and a homemade dual-glow luciferase assay. This protocol rapidly generates direct side-by-side functional data for thousands of genes and is easily modifiable to target any gene of interest.

We present a rapid and inexpensive high-throughput screening protocol to identify transcriptional regulators of alpha-synuclein, a gene associated with ...

Immunodetection of Outer Membrane Proteins by Flow Cytometry of Isolated Mitochondria

Methods to detect and monitor mitochondrial outer membrane protein components in animal tissues are vital to study mitochondrial physiology and pathophysiology. This protocol describes a technique where mitochondria isolated from rodent tissue are immunolabeled and analyzed by flow cytometry. Mitochondria are isolated from rodent spinal cords and subjected to a rapid enrichment step so as to remove myelin, a major contaminant of mitochondrial fractions prepared from nervous tissue. Isolated mitochondria are then labeled with an antibody of choice and a fluorescently conjugated secondary antibody. Analysis by flow cytometry verifies the relative purity of mitochondrial preparations by staining with a mitochondrial specific dye, followed by detection and quantification of immunolabeled protein. This technique is rapid, quantifiable and high-throughput, allowing for the analysis of hundreds of thousands of mitochondria per sample. It is applicable to assess novel proteins at the mitochondrial surface under normal physiological conditions as well as the proteins that may become mislocalized to this organelle during pathology. Importantly, this method can be coupled to fluorescent indicator dyes to report on certain activities of mitochondrial subpopulations and is feasible for mitochondria from the central nervous system (brain and spinal cord) as well as liver.

Described here is a method to detect and quantify mitochondrial outer membrane proteins by immunolabeling of mitochondria isolated from rodent tissue and analysis by flow cytometry. This method can be extended to assess functional aspects of mitochondrial subpopulations.