We would like to thank all members of the Chipuk Laboratory for their colleagueship and support. In addition, we would like to give appreciation to Tomomi Kuwana and Donald Newmeyer for developing the experimental foundation for this work. This work was supported by: NIH CA157740 (to J.E.C.), and a pilot project from NIH P20AA017067 (to J.E.C.). This work was also supported in part by a Research Grant 5-FY11-74 from the March of Dimes Foundation (to J.E.C.).

1. Combine Lipids and Create a Lipid Film
<ol>
	<li>Working in a fume hood with minimal lighting, chloroform-solubilized lipids are combined into a 1.5 ml amber glass vial using a chloroform-rinsed Hamilton gas-tight glass syringe. This combination will yield approximately 4 mg of lipid (Figure 1A).
	<ul>
		<li>Cardiolipin 10 mg/ml (CL) - 63 &mu;l</li>
		<li>Phosphatidylcholine 10 mg/ml (PC) - 84 &mu;l</li>
		<li>Phosphatidylethanoloamine 10 mg/ml (PE) - 125 &mu;l</li>
		<li>Phosphatidylinositol 10 mg/ml (PI) - 59.5 &mu;l</li>
		<li>Phosphatidylserine 10 mg/ml (PS) - 65 &mu;l</li>
	</ul>
	</li>
	<li>To remove the chloroform, apply a gentle stream of an inert gas (e.g., nitrogen or argon) into the vial. A lipid film will become visible on the bottom and sides of the vial when dry, which typically takes 2 - 3 hours (Figure 1A).</li>
</ol>
2. Prepare Liposome Solution
<ol>
	<li>In 1.7 ml microcentrifuge tubes, prepare 500 &mu;l of 12.5 mM ANTS in LUV buffer (0.2 mM EDTA, 10 mM HEPES [pH 7], 200 mM KCl, 5 mM MgCl2 ) and 500 &mu;l of 45 mM DPX in LUV buffer (Figure 1B).</li>
	<li>Combine the ANTS and DPX solutions, vortex for 1 min, and sonicate for 1 min at room temperature. Sonication is done in an ultrasonic water bath (e.g., Branson 3510MTH, no setting, just 'on'). This ensures the solutions are entirely solubilized and homogeneous.</li>
	<li>Add the ANTS/DPX solution to the lipid film, and seal the vial with parafilm (Figure 1B).</li>
	<li>Sonicate the lipid film in the above water bath for 5 min. The solution will appear milky after sonication. Ensure the lipid film is completely solubilized as small lipid aggregates will negatively impact on proceeding steps. Also, check the vial's sides and bottom to ensure there is no remaining lipid film (Figure 1B).</li>
</ol>
3. Extrusion and Purification of Liposomes
<ol>
	<li>Assemble the extruder according to the manufacturer's guidelines using LUV buffer to moisten the filter supports and polycarbonate membrane. Tighten the unit's casing so that there will be no loss of volume during the extrusion, but not overly tight as to rupture the membrane. It's also wise to test the extruder with LUV buffer to ensure there are no leaks or problems.</li>
	<li>Start with the lipid solution in the right syringe, extrude the lipid solution 31 times, and the LUVs will finish in the left syringe; this ensures all the lipids have passed through the membrane at least once. The first few times may required a little more pressure to pass through the membrane. If the solution suddenly passes through the membrane quickly, check the membrane for rupture and replace, if necessary (Figure 1C).</li>
	<li>Set-up a 10 ml Sepharose S-500 gravity flow column (1.0 cm x 20 cm, 10 ml bead volume). Wash with 3 column volumes of LUV buffer, allow the column flow to stop, and cap. Since this step takes a while, it is preferred to start the washing when the lipids are combined to dry. Just keep the column capped until you are ready to use it.</li>
	<li>Add the extruded LUVs to the column in one step, avoid disrupting the bead interface. Allow the buffer to run out of the column, the flow will stop, and plug. Gently fill the column with LUV buffer. Again, use caution when adding buffer so that the beads are not disturbed (Figure 1C).</li>
	<li>Remove the plug and collect 6 x 1 ml fractions. The LUVs normally elute in fractions 3 and 4, and can be identified by a cloudy appearance. Once the fractions are collected, wash the column with 5 volumes of water, and 2 volumes of 20% ethanol. Store the column at 4 &deg;C (Figure 1C).</li>
	<li>Store the fractions at 4 &deg;C in the dark until tested.</li>
</ol>
4. Test Column Fractions
<ol>
	<li>The column fractions are now tested for fluorescence comparing LUV buffer and LUV buffer supplemented with 0.5% CHAPS. Fractions that increase in fluorescence due to the presence CHAPS contain LUVs. Pipette 100 &mu;l of LUV buffer or LUV buffer + 0.5% CHAPS into six wells each, and add 5 &mu;l of the appropriate fraction, mix. Use opaque, black, untreated, 96 well plates for all LUV studies (Figure 1D).</li>
	<li>Read the plate at 37 &deg;C with the following parameters: Excitation wavelength: 355 nm; Emission wavelength: 520 nm; Gain (voltage): 125; Optics position: Top; Read height: 5.5 mm. This set-up is for the BioTek H1 Synergy using Gen 5 2.0 software (Figure 1D).</li>
	<li>Fractions 3 and 4 generally show a 5 - 10 fold increase in fluorescence. Once analyzed, combine the LUV-containing fractions and store at 4 &deg;C in the dark; approximately 2 ml of ~ 2 mM lipid will be purified (Figure 1D).</li>
</ol>
5. BCL-2 Family Studies Using LUVs
<ol>
	<li>BCL-2 proteins are tested for their ability to directly induce LUV permeabilization, and for the regulation of permeabilization activity. LUV assays are normally performed in 100 &mu;l volumes, at 37 &deg;C, and can be analyzed as either a single time point after 30 - 60 min (Figures 2A-C, 3A-B), or via kinetic studies every 1 - 2 min for up to 1 hr (Figure 2D). A representative assay design describing components, stock/final concentrations, and order of addition, is provided (Table 1). It is also advisable to have the microplate reader gently shake the plate prior to each reading. The first few min often give sporadic readings (likely due to temperature changes, mixing, etc...), so we generally consider data points after 2 - 3 readings in a kinetic study. It is also not uncommon to generate large error bars for some of the conditions due to variability in LUV batches, and the efficiency of protein interactions at borderline concentrations of activation or inhibition.</li>
	<li>BCL-2 family protein stocks should be diluted into LUV buffer at a concentration of 100x, and necessary titrations of BAX, BID, and all other proteins are performed to find the lowest concentrations necessary to detect minimal BAX-dependent LUV permeabilization in the absence of BID, and significant synergy between BAX and BID. Furthermore, all other proteins require titrations to reduce protein waste, and minimize the concentrations of subsequent protein additions.</li>
</ol>
6. Representative Results
There are a myriad of options to witness and examine BCL-2 family dependent regulation of LUV permeabilization. To generate a LUV permeabilization positive control, we use detergent-activated BAX12. The detergent, n-octyl-&beta;-D-glucoside (OG), artificially triggers BAX activation (for 100 &mu;l of 2.3 &mu;M OG-BAX: 5 &mu;g BAX + 0.7% OG in LUV buffer, incubate for 60 min at 4 &deg;C, aliquot and store at -80 &deg;C), and therefore OG-BAX (10 - 100 nM) can be used as a reliable positive control. Of note, OG concentrations below 0.025% do not directly affect LUV permeabilization. A 0.5% CHAPS treatment is used to determine the maximum amount of LUV fluorescence per assay, and this value sets the 100% value. All treatments should be set-up in triplicate. An example LUV control assay set-up is shown in Table 1, and corresponding data are presented in Figure 2A.
A necessary starting point for studying LUV permeabilization with BCL-2 family proteins is to establish the concentrations of BAX and BID to promote optimal release. As individual protein preparations will demonstrate different background permeabilization activities, it is advisable to titrate before setting up more sophisticated analyses. BAX (50 - 200 nM) and BID (0 - 10 nM) must be titrated for minimal background release, and optimal synergy, as shown in Figures 2B-C, respectively. Various forms of BID protein can be purchased from R&amp;D Systems; alternatively, the BID BH3 domain peptide is available from Anaspec. There is no commercial form of BAX that is appropriate for LUV assays, therefore it must be expressed and purified using published methods3,13.
Once BAX and BID concentrations and synergy are established, a common assay is to determine the influence of anti-apoptotic proteins on BAX and BID-dependent LUV permeabilization. BCL-xL is an anti-apoptotic BCL-2 member that is commercially available, and in a dose-dependent manner, inhibits the activity of BAX and BID usually in the range of 5 - 10 fold molar excess of BAX or BID (Figure 3A). This inhibitory effect can be reversed by the addition of subsequent BH3-only proteins (2 - 5 fold molar excess to BCL-xL) or BH3 domain peptides (5 - 10 fold molar excess to BCL-xL), which bind to BCL-xL, thus allowing for BAX and BID to synergize and permeabilize the LUVs (Figure 3B). In Figure 3B, we show the effect of full-length recombinant human PUMA&beta;, and the PUMA BH3 domain peptide is functionally similar (data not shown)8. This experiment is commonly referred to as a 'de-repression assay', as the PUMA protein relieves the inhibitory effect of BCL-xL on BAX and BID. Ideally, the de-repressing BH3-only protein/peptide addition should have minimal effects on BAX activation14.
<img alt="Figure 1" fo:content-width="6.5in" fo:src="/files/ftp_upload/4291/4291fig1highres.jpg" src="/files/ftp_upload/4291/4291fig1.jpg" /> 
Figure 1. (A) Chloroform solubilized lipids are combined, dried to create a lipid film, and capped. (B) ANTS and DPX solutions are prepared, combined, and sonicated to ensure complete solubilization and homogenization. The solution is added to the lipid film, the vial is sealed with parafilm, and sonicated for 5 min in a water bath. The solution will now appear milky. (C) The sonicated lipid solution is loaded into the mini-extruder, processed 31 times, and added to a prepared Sepharose S-500 column for subsequent purification. (D) Fractions obtained after size exclusion chromatography are analyzed by comparing the relative fluorescent units (RFUs) in LUV buffer and LUV buffer + 0.5% CHAPS. The "LUV buffer" reading indicates background fluorescence with intact LUVs; the "LUV buffer + 0.5% CHAPS" condition will show lysis induced gain of fluorescence, indicative of LUVs. Fractions 3 and 4 usually contain the LUVs, and are combined after analysis. Fractions 5 and 6 contain unincorporated ANTS (i.e., not within the LUVs following extrusion), which show high fluorescence independent of CHAPS. <a href="https://www.jove.com/files/ftp_upload/4291/4291fig1large.jpg" target="_blank"> Click here to view larger figure</a>.
<img alt="Figure 2" fo:content-width="5.45in" fo:src="/files/ftp_upload/4291/4291fig2highres.jpg" src="/files/ftp_upload/4291/4291fig2.jpg" /> 
Figure 2. BCL-2 family control experiments and BAX/BID titrations using LUVs. (A) Control experiments. LUVs were treated with LUV buffer alone, 0.025% octylglucoside "OG", 50 nM BAX, or 50 nM OG-BAX. LUVs were incubated for 60 min at 37 &deg;C. The percent (%) permeabilization is determined by subtracting the fluorescence in the buffer treatment from all the samples, 100% permeabilization is determined by a 0.5% CHAPS solubilized sample; divide the experimental values by the CHAPS value to obtain % LUV permeabilization. OG-BAX is detergent-activated BAX, which is a reliable positive control for LUV permeabilization. (B) Determine the optimal BAX concentration. LUVs were treated with increasing concentrations of BAX (50 - 200 nM) in the presence of 10 nM C8-BID "BID", or buffer alone. (C) Determine the optimal BID concentration. LUVs were treated with increasing concentrations of BID (0 - 10 nM) in the presence of 100 nM BAX. (D) An example of a LUV permeabilization kinetic dataset. LUVs were treated with BAX (100 nM) &plusmn; BID (10 nM). Readings were performed every two min for 30 min, with two sec of mild plate shaking prior to each read. The first three readings (0, 2, &amp; 4 min) are not included in the graph. The minimum and maximum relative fluorescent units (RFU) were determined as described in section 4.3. The error bars in Figures 2A-C represent the standard deviation from triplicate data. Data in 2D represent a duplicate assay.
<img alt="Figure 3" fo:content-width="4in" fo:src="/files/ftp_upload/4291/4291fig3highres.jpg" src="/files/ftp_upload/4291/4291fig3.jpg" /> 
Figure 3. Examples of common BCL-2 family interactions explored using LUVs. (A) BAX and BID mediated LUV permeabilization is inhibited by BCL-xL. LUVs were incubated with 100 nM BAX, 10 nM BID, and increasing concentrations of BCL-xL (0 - 500 nM). LUVs were incubated for 60 min at 37 &deg;C. (B) PUMA de-represses BCL-xL to promote BAX and BID function. LUVs were incubated with combinations of 100 nM BAX, 10 nM BID, 500 nM BCL-xL, and (0 - 2.5 &mu;M) PUMA for 60 min at 37 &deg;C. PUMA was used at 2.5 &mu;M with BAX alone. The error bars represent the standard deviation from triplicate data.
<table border="1">
	<tbody>
		<tr>
			<td>Order of addition:</td>
			<td>1</td>
			<td>2</td>
			<td>3</td>
			<td>4</td>
			<td>5</td>
			<td>6</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>CONDITIONS</td>
			<td>LUV 
			Buffer</td>
			<td>OG 
			(0.7%)</td>
			<td>BAX 
			(2.3 &mu;M)</td>
			<td>OG-BAX 
			(2.3 &mu;M)</td>
			<td>CHAPS 
			(10%)</td>
			<td>LUVs 
			(2 mM)</td>
			<td>TOTAL</td>
		</tr>
		<tr>
			<td>Buffer</td>
			<td>95 &mu;l</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>5 &mu;l</td>
			<td>100 &mu;l</td>
		</tr>
		<tr>
			<td>OG (0.015%)</td>
			<td>92.83 &mu;l</td>
			<td>2.17 &mu;l</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>5 &mu;l</td>
			<td>100 &mu;l</td>
		</tr>
		<tr>
			<td>BAX (50 nM)</td>
			<td>92.83 &mu;l</td>
			<td>-</td>
			<td>2.17 &mu;l</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>5 &mu;l</td>
			<td>100 &mu;l</td>
		</tr>
		<tr>
			<td>OG-BAX (50 nM)</td>
			<td>92.83 &mu;l</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>2.17 &mu;l</td>
			<td>&nbsp;</td>
			<td>5 &mu;l</td>
			<td>100 &mu;l</td>
		</tr>
		<tr>
			<td>CHAPS (0.5%)</td>
			<td>90 &mu;l</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>5 &mu;l</td>
			<td>5 &mu;l</td>
			<td>100 &mu;l</td>
		</tr>
	</tbody>
</table>
Table 1. A control assay example to determine the effect of OG-BAX on LUV permeabilization.

<ol>
	<li>Chipuk, J. E., Moldoveanu, T., Llambi, F., Parsons, M. J., Green, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+BCL-2+family+reunion.">The BCL-2 family reunion.</a> Mol. Cell. 37, 299-310 (2010).</li><li>Green, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Apoptotic+pathways:+ten+minutes+to+dead.">Apoptotic pathways: ten minutes to dead.</a> Cell. 121, 671-674 (2005).</li><li>Gavathiotis, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=BAX+activation+is+initiated+at+a+novel+interaction+site.">BAX activation is initiated at a novel interaction site.</a> Nature. 455, 1076-1081 (2008).</li><li>Dewson, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=To+trigger+apoptosis,+Bak+exposes+its+BH3+domain+and+homodimerizes+via+BH3:groove+interactions.">To trigger apoptosis, Bak exposes its BH3 domain and homodimerizes via BH3:groove interactions.</a> Mol. Cell. 30, 369-380 (2008).</li><li>Dewson, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bak+activation+for+apoptosis+involves+oligomerization+of+dimers+via+their+alpha6+helices.">Bak activation for apoptosis involves oligomerization of dimers via their alpha6 helices.</a> Mol. Cell. 36, 696-703 (2009).</li><li>Gavathiotis, E., Reyna, D. E., Davis, M. L., Bird, G. H., Walensky, L. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=BH3-triggered+structural+reorganization+drives+the+activation+of+proapoptotic+BAX.">BH3-triggered structural reorganization drives the activation of proapoptotic BAX.</a> Mol. Cell. 40, 481-492 (2010).</li><li>Kuwana, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=BH3+domains+of+BH3-only+proteins+differentially+regulate+Bax-mediated+mitochondrial+membrane+permeabilization+both+directly+and+indirectly.">BH3 domains of BH3-only proteins differentially regulate Bax-mediated mitochondrial membrane permeabilization both directly and indirectly.</a> Mol. Cell. 17, 525-535 (2005).</li><li>Chipuk, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanism+of+apoptosis+induction+by+inhibition+of+the+anti-apoptotic+BCL-2+proteins.">Mechanism of apoptosis induction by inhibition of the anti-apoptotic BCL-2 proteins.</a> Proc. Natl. Acad. Sci. U.S.A. 105, 20327-20332 (2008).</li><li>Letai, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distinct+BH3+domains+either+sensitize+or+activate+mitochondrial+apoptosis,+serving+as+prototype+cancer+therapeutics.">Distinct BH3 domains either sensitize or activate mitochondrial apoptosis, serving as prototype cancer therapeutics.</a> Cancer Cell. 2, 183-192 (2002).</li><li>Kuwana, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=lipids+cooperate+to+form+supramolecular+openings+in+the+outer+mitochondrial+membrane.">lipids cooperate to form supramolecular openings in the outer mitochondrial membrane.</a> Cell. 111, 331-342 (2002).</li><li>Lovell, J. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Membrane+binding+by+tBid+initiates+an+ordered+series+of+events+culminating+in+membrane+permeabilization+by+Bax.">Membrane binding by tBid initiates an ordered series of events culminating in membrane permeabilization by Bax.</a> Cell. 135, 1074-1084 (2008).</li><li>Hsu, Y. T., Youle, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nonionic+detergents+induce+dimerization+among+members+of+the+Bcl-2+family.">Nonionic detergents induce dimerization among members of the Bcl-2 family.</a> J Biol Chem. 272, 13829-13834 (1997).</li><li>Suzuki, M., Youle, R. J., Tjandra, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structure+of+Bax:+coregulation+of+dimer+formation+and+intracellular+localization.">Structure of Bax: coregulation of dimer formation and intracellular localization.</a> Cell. 103, 645-654 (2000).</li><li>Du, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=BH3+domains+other+than+Bim+and+Bid+can+directly+activate+Bax/Bak.">BH3 domains other than Bim and Bid can directly activate Bax/Bak.</a> J Biol Chem. 286, 491-501 (2011).</li></ol>

No conflicts of interest declared.

The described method for generating LUVs enables a rapid and efficient means to test the function of various BCL-2 family proteins, peptides, and related reagents in a biochemically-defined membrane environment similar to the OMM. If using end point values to determine LUV permeabilization, multiple plates can be set-up to analyze hundreds of conditions within a single day. We find that the limiting reagents in these assays tend to be the quality and quantity of recombinant proteins, so dedicating sufficient time and res...

<table border="1">
	<tbody>
		<tr>
			<td>Name of the reagent</td>
			<td>Company</td>
			<td>Catalogue number</td>
			<td>Comments (optional)</td>
		</tr>
		<tr>
			<td>1,2-Dioleoyl-sn-Glycero-3-Phosphoethanolamine "PE"</td>
			<td>Avanti</td>
			<td>850725C</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>L-&alpha;-Phosphatidylcholine (Egg, Chicken) "PC"</td>
			<td>Avanti</td>
			<td>840051C</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>L-&alpha;-Phosphatidylinositol (Liver, Bovine) "PI"</td>
			<td>Avanti</td>
			<td>840042C</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>L-&alpha;-Phosphatidylserine (Brain, Porcine) "PS"</td>
			<td>Avanti</td>
			<td>8400320</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Cardiolipin (Heart, Bovine - Sodium Salt) "CL"</td>
			<td>Avanti</td>
			<td>840012C</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Mini-extruder set</td>
			<td>Avanti</td>
			<td>610023</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>PC membrane 0.2 &mu;M</td>
			<td>Avanti</td>
			<td>610006</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Costar black 96 well plate</td>
			<td>Fisher Scientific</td>
			<td>07-200-590</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Caspase-8 cleaved human BID</td>
			<td>R&amp;D Systems</td>
			<td>882-B8-050</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Human BCL-xL (minus C-terminus)</td>
			<td>R&amp;D Systems</td>
			<td>894-BX-050</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>BID/PUMA BH3 domain peptides</td>
			<td>Anaspec</td>
			<td>61711/62404</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Synergy H1 hybrid multi-mode microplate reader</td>
			<td>BioTek</td>
			<td>None</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
</table>

examining bcl-2 family function with large unilamellar vesicles

The BCL-2 (B cell CLL/Lymphoma) family is comprised of approximately twenty proteins that collaborate to either maintain cell survival or initiate apoptosis1. Following cellular stress (e.g., DNA damage), the pro-apoptotic BCL-2 family effectors BAK (BCL-2 antagonistic killer 1) and/or BAX (BCL-2 associated X protein) become activated and compromise the integrity of the outer mitochondrial membrane (OMM), though the process referred to as mitochondrial outer membrane permeabilization (MOMP)1. After MOMP occurs, pro-apoptotic proteins (e.g., cytochrome c) gain access to the cytoplasm, promote caspase activation, and apoptosis rapidly ensues2.
In order for BAK/BAX to induce MOMP, they require transient interactions with members of another pro-apoptotic subset of the BCL-2 family, the BCL-2 homology domain 3 (BH3)-only proteins, such as BID (BH3-interacting domain agonist)3-6. Anti-apoptotic BCL-2 family proteins (e.g., BCL-2 related gene, long isoform, BCL-xL; myeloid cell leukemia 1, MCL-1) regulate cellular survival by tightly controlling the interactions between BAK/BAX and the BH3-only proteins capable of directly inducing BAK/BAX activation7,8. In addition, anti-apoptotic BCL-2 protein availability is also dictated by sensitizer/de-repressor BH3-only proteins, such as BAD (BCL-2 antagonist of cell death) or PUMA (p53 upregulated modulator of apoptosis), which bind and inhibit anti-apoptotic members7,9. As most of the anti-apoptotic BCL-2 repertoire is localized to the OMM, the cellular decision to maintain survival or induce MOMP is dictated by multiple BCL-2 family interactions at this membrane.
 Large unilamellar vesicles (LUVs) are a biochemical model to explore relationships between BCL-2 family interactions and membrane permeabilization10. LUVs are comprised of defined lipids that are assembled in ratios identified in lipid composition studies from solvent extracted Xenopus mitochondria (46.5% phosphatidylcholine, 28.5% phosphatidylethanoloamine, 9% phosphatidylinositol, 9% phosphatidylserine, and 7% cardiolipin)10. This is a convenient model system to directly explore BCL-2 family function because the protein and lipid components are completely defined and tractable, which is not always the case with primary mitochondria. While cardiolipin is not usually this high throughout the OMM, this model does faithfully mimic the OMM to promote BCL-2 family function. Furthermore, a more recent modification of the above protocol allows for kinetic analyses of protein interactions and real-time measurements of membrane permeabilization, which is based on LUVs containing a polyanionic dye (ANTS: 8-aminonaphthalene-1,3,6-trisulfonic acid) and cationic quencher (DPX: p-xylene-bis-pyridinium bromide)11. As the LUVs permeabilize, ANTS and DPX diffuse apart, and a gain in fluorescence is detected. Here, commonly used recombinant BCL-2 family protein combinations and controls using the LUVs containing ANTS/DPX are described.

Watch this Scientific Journal Video about Examining BCL-2 Family Function with Large Unilamellar Vesicles at JoVE.com

Examining BCL-2 Family Function with Large Unilamellar Vesicles

Biochemically-defined large unilamellar vesicles (LUVs) are a convenient model system to analyze BCL-2 family interactions with immediate implications in better understanding the mitochondrial pathway of apoptosis. A method to produce LUVs, along with standard BCL-2 family protein combinations and controls to examine LUV permeabilization, are presented.

The BCL-2 (B cell CLL/Lymphoma) family is comprised of approximately twenty proteins that collaborate to either maintain cell survival or initiate ...

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7.8K Views.  Mount Sinai School of Medicine. Biochemically-defined large unilamellar vesicles (LUVs) are a convenient model system to analyze BCL-2 family interactions with immediate implications in better understanding the mitochondrial pathway of apoptosis. A method to produce LUVs, along with standard BCL-2 family protein combinations and controls to examine LUV permeabilization, are presented.

Video: Examining BCL-2 Family Function with Large Unilamellar Vesicles

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2, ion pumps3, and transporters4. Recent developments have combined site-specific fluorophore labeling alongside TEVC to 
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8. 
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4. Finally, recent developments 
have also enabled the manipulation of select internal ions following co-expression with a second protein9.
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (&lt;5%)3 upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of 
the membrane protein in order to correlate the conformational dynamics to the function of the protein10. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

We will describe a method which measures the kinetics of ion transport of membrane proteins alongside site-specific analysis of conformational changes using fluorescence on single cells. This technique is adaptable to ion channels, transporters and ion pumps and can be utilized to determine distance constraints between protein subunits.

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane ...

Measuring Peptide Translocation into Large Unilamellar Vesicles

There is an active interest in peptides that readily cross cell membranes without the assistance of cell membrane receptors1. Many of these are referred to as cell-penetrating peptides, which are frequently noted for their potential as drug delivery vectors1-3. Moreover, there is increasing interest in antimicrobial peptides that operate via non-membrane lytic mechanisms4,5, particularly those that cross bacterial membranes without causing cell lysis and kill cells by interfering with intracellular processes6,7. In fact, authors have increasingly pointed out the relationship between cell-penetrating and antimicrobial peptides1,8. A firm understanding of the process of membrane translocation and the relationship between peptide structure and its ability to translocate requires effective, reproducible assays for translocation. Several groups have proposed methods to measure translocation into large unilamellar lipid vesicles (LUVs)9-13. LUVs serve as useful models for bacterial and eukaryotic cell membranes and are frequently used in peptide fluorescent studies14,15. Here, we describe our application of the method first developed by Matsuzaki and co-workers to consider antimicrobial peptides, such as magainin and buforin II16,17. In addition to providing our protocol for this method, we also present a straightforward approach to data analysis that quantifies translocation ability using this assay. The advantages of this translocation assay compared to others are that it has the potential to provide information about the rate of membrane translocation and does not require the addition of a fluorescent label, which can alter peptide properties18, to tryptophan-containing peptides. Briefly, translocation ability into lipid vesicles is measured as a function of the Foster Resonance Energy Transfer (FRET) between native tryptophan residues and dansyl phosphatidylethanolamine when proteins are associated with the external LUV membrane (Figure 1). Cell-penetrating peptides are cleaved as they encounter uninhibited trypsin encapsulated with the LUVs, leading to disassociation from the LUV membrane and a drop in FRET signal. The drop in FRET signal observed for a translocating peptide is significantly greater than that observed for the same peptide when the LUVs contain both trypsin and trypsin inhibitor, or when a peptide that does not spontaneously cross lipid membranes is exposed to trypsin-containing LUVs. This change in fluorescence provides a direct quantification of peptide translocation over time.

This protocol details a method for the quantitative measure of peptide translocation into large unilamellar lipid vesicles. This method also provides information about the rate of membrane translocation and can be used to identify peptides that efficiently and spontaneously cross lipid bilayers.

There is an active interest in peptides that readily cross cell membranes without the assistance of cell membrane receptors1.  Many of these are referred ...

Cryosectioning Yeast Communities for Examining Fluorescence Patterns

Microbes typically live in communities. The spatial organization of cells within a community is believed to impact the survival and function of the community1. Optical sectioning techniques, including confocal and two-photon microscopy, have proven useful for observing spatial organization of bacterial and archaeal communities2,3. A combination of confocal imaging and physical sectioning of yeast colonies has revealed internal organization of cells4. However, direct optical sectioning using confocal or two-photon microscopy has been only able to reach a few cell layers deep into yeast colonies. This limitation is likely because of strong scattering of light from yeast cells4.
Here, we present a method based on fixing and cryosectioning to obtain spatial distribution of fluorescent cells within Saccharomyces cerevisiae communities. We use methanol as the fixative agent to preserve the spatial distribution of cells. Fixed communities are infiltrated with OCT compound, frozen, and cryosectioned in a cryostat. Fluorescence imaging of the sections reveals the internal organization of fluorescent cells within the community.
Examples of yeast communities consisting of strains expressing red and green fluorescent proteins demonstrate the potentials of the cryosectioning method to reveal the spatial distribution of fluorescent cells as well as that of gene expression within yeast colonies2,3. Even though our focus has been on Saccharomyces cerevisiae communities, the same method can potentially be applied to examine other microbial communities.

We present a protocol for freezing and cryosectioning yeast communities to observe internal patterns of fluorescent cells. The method relies on methanol-fixing and OCT-embedding to preserve the spatial distribution of cells without inactivating fluorescent proteins within a community.

Microbes typically live  in communities. The spatial organization  of cells within a community is believed to impact the survival and function of the  ...

Large-scale Gene Knockdown in C. elegans Using dsRNA Feeding Libraries to Generate Robust Loss-of-function Phenotypes

RNA interference by feeding worms bacteria expressing dsRNAs has been a useful tool to assess gene function in C. elegans. While this strategy works well when a small number of genes are targeted for knockdown, large scale feeding screens show variable knockdown efficiencies, which limits their utility. We have deconstructed previously published RNAi knockdown protocols and found that the primary source of the reduced knockdown can be attributed to the loss of dsRNA-encoding plasmids from the bacteria fed to the animals. Based on these observations, we have developed a dsRNA feeding protocol that greatly reduces or eliminates plasmid loss to achieve efficient, high throughput knockdown. We demonstrate that this protocol will produce robust, reproducible knock down of C. elegans genes in multiple tissue types, including neurons, and will permit efficient knockdown in large scale screens. This protocol uses a commercially available dsRNA feeding library and describes all steps needed to duplicate the library and perform dsRNA screens. The protocol does not require the use of any sophisticated equipment, and can therefore be performed by any C. elegans lab.

Large-scale Gene Knockdown in C. elegans Using dsRNA Feeding Libraries to Generate Robust Loss-of-function Phenotypes

While dsRNA feeding in C. elegans is a powerful tool to assess gene function, current protocols for large scale feeding screens result in variable knockdown efficiencies. We describe an improved protocol for performing large scale RNAi feeding screens that results in highly efficacious and reproducible knockdown of gene expression.

RNA interference by feeding worms bacteria expressing dsRNAs has been a useful tool to assess gene function in C. elegans. While this strategy works well ...

Phenotyping Mouse Pulmonary Function In Vivo with the Lung Diffusing Capacity

The mouse is now the primary animal used to model a variety of lung diseases. To study the mechanisms that underlie such pathologies, phenotypic methods are needed that can quantify the pathologic changes. Furthermore, to provide translational relevance to the mouse models, such measurements should be tests that can easily be done in both humans and mice. Unfortunately, in the present literature few phenotypic measurements of lung function have direct application to humans. One exception is the diffusing capacity for carbon monoxide, which is a measurement that is routinely done in humans. In the present report, we describe a means to quickly and simply measure this diffusing capacity in mice. The procedure involves brief lung inflation with tracer gases in an anesthetized mouse, followed by a 1 min gas analysis time. We have tested the ability of this method to detect several lung pathologies, including emphysema, fibrosis, acute lung injury, and influenza and fungal lung infections, as well as monitoring lung maturation in young pups. Results show significant decreases in all the lung pathologies, as well as an increase in the diffusing capacity with lung maturation. This measurement of lung diffusing capacity thus provides a pulmonary function test that has broad application with its ability to detect phenotypic structural changes with most of the existing pathologic lung models.

Phenotyping Mouse Pulmonary Function In Vivo with the Lung Diffusing Capacity

We describe a means to quickly and simply measure the lung diffusing capacity in mice and show that it is sufficiently sensitive to phenotype changes in multiple common lung pathologies. This metric thus brings direct translational relevance to the mouse models, since diffusing capacity is also easily measured in humans.

The mouse is now the primary animal used to model a variety of lung diseases. To study the mechanisms that underlie such pathologies, phenotypic methods ...

Reconstitution of a Transmembrane Protein, the Voltage-gated Ion Channel, KvAP, into Giant Unilamellar Vesicles for Microscopy and Patch Clamp Studies

Giant Unilamellar Vesicles (GUVs) are a popular biomimetic system for studying membrane associated phenomena. However, commonly used protocols to grow GUVs must be modified in order to form GUVs containing functional transmembrane proteins. This article describes two dehydration-rehydration methods &#8212; electroformation and gel-assisted swelling &#8212; to form GUVs containing the voltage-gated potassium channel, KvAP. In both methods, a solution of protein-containing small unilamellar vesicles is partially dehydrated to form a stack of membranes, which is then allowed to swell in a rehydration buffer. For the electroformation method, the film is deposited on platinum electrodes so that an AC field can be applied during film rehydration. In contrast, the gel-assisted swelling method uses an agarose gel substrate to enhance film rehydration. Both methods can produce GUVs in low (e.g., 5 mM) and physiological (e.g., 100 mM) salt concentrations. The resulting GUVs are characterized via fluorescence microscopy, and the function of reconstituted channels measured using the inside-out patch-clamp configuration. While swelling in the presence of an alternating electric field (electroformation) gives a high yield of defect-free GUVs, the gel-assisted swelling method produces a more homogeneous protein distribution and requires no special equipment.

The reconstitution of the transmembrane protein, KvAP, into giant unilamellar vesicles (GUVs) is demonstrated for two dehydration-rehydration methods &#8212; electroformation, and gel-assisted swelling. In both methods, small unilamellar vesicles containing the protein are fused together to form GUVs that can then be studied by fluorescence microscopy and patch-clamp electrophysiology.

Giant Unilamellar Vesicles (GUVs) are a popular biomimetic system for studying membrane associated phenomena. However, commonly used protocols to grow ...

Visualization of Surface-tethered Large DNA Molecules with a Fluorescent Protein DNA Binding Peptide

Large DNA molecules tethered on the functionalized glass surface have been utilized in polymer physics and biochemistry particularly for investigating interactions between DNA and its binding proteins. Here, we report a method that uses fluorescent microscopy for visualizing large DNA molecules tethered on the surface. First, glass coverslips are biotinylated and passivated by coating with biotinylated polyethylene glycol, which specifically binds biotinylated DNA via avidin protein linkers and significantly reduces undesirable binding from non-specific interactions of proteins or DNA molecules on the surface. Second, the DNA molecules are biotinylated by two different methods depending on their terminals. The blunt ended DNA is tagged with biotinylated dUTP at its 3' hydroxyl terminus, by terminal transferase, while the sticky ended DNA is hybridized with biotinylated complimentary oligonucleotides by DNA ligase. Finally, a microfluidic shear flow makes single DNA molecules stretch to their full contour lengths after being stained with fluorescent protein-DNA binding peptide (FP-DBP).

We present an approach for visualizing fluorescent protein DNA binding peptide (FP-DBP)-stained large DNA molecules tethered on the polyethylene glycol (PEG) and avidin-coated glass surface and stretched with microfluidic shear flows.

Large DNA molecules tethered on the functionalized glass surface have been utilized in polymer physics and biochemistry particularly for investigating ...

Pulling Membrane Nanotubes from Giant Unilamellar Vesicles

The reshaping of the cell membrane is an integral part of many cellular phenomena, such as endocytosis, trafficking, the formation of filopodia, etc. Many different proteins associate with curved membranes because of their ability to sense or induce membrane curvature. Typically, these processes involve a multitude of proteins making them too complex to study quantitatively in the cell. We describe a protocol to reconstitute a curved membrane in vitro, mimicking a curved cellular structure, such as the endocytic neck. A giant unilamellar vesicle (GUV) is used as a model of a cell membrane, whose internal pressure and surface tension are controlled with micropipette aspiration. Applying a point pulling force on the GUV using optical tweezers creates a nanotube of high curvature connected to a flat membrane. This method has traditionally been used to measure the fundamental mechanical properties of lipid membranes, such as bending rigidity. In recent years, it has been expanded to study how proteins interact with membrane curvature and the way they affect the shape and the mechanics of membranes. A system combining micromanipulation, microinjection, optical tweezers, and confocal microscopy allows measurement of membrane curvature, membrane tension, and the surface density of proteins, concurrently. From these measurements, many important mechanical and morphological properties of the protein-membrane system can be inferred. In addition, we lay out a protocol of creating GUVs in the presence of physiological salt concentration, and a method of quantifying the surface density of proteins on the membrane from fluorescence intensities of labeled proteins and lipids.

Many proteins in the cell sense and induce membrane curvature. We describe a method to pull membrane nanotubes from lipid vesicles to study the interaction of proteins or any curvature-active molecule with curved membranes in vitro.

The reshaping of the cell membrane is an integral part of many cellular phenomena, such as endocytosis, trafficking, the formation of filopodia, etc. Many ...

A Bacterial Oral Feeding Assay with Antibiotic-Treated Mosquitoes

The mosquito midgut harbors a highly dynamic microbiome that affects the host metabolism, reproduction, fitness, and vector competence. Studies have been conducted to investigate the effect of gut microbes as a whole; however, different microbes could exert distinct effects toward the host. This article provides the methodology to study the effect of each specific mosquito gut microbe and the potential mechanism.
This protocol contains two parts. The first part introduces how to dissect the mosquito midgut, isolate cultivable bacteria colonies, and identify bacteria species. The second part provides the procedure to generate antibiotic-treated mosquitoes and reintroduce one specific bacteria species.

This article presents a protocol to investigate the effect of individual mosquito gut bacteria, including isolation and identification of mosquito midgut cultivable microbes, antibiotic depletion of mosquito gut bacteria, and reintroduce one specific bacteria species.

The mosquito midgut harbors a highly dynamic microbiome that affects the host metabolism, reproduction, fitness, and vector competence. Studies have been ...

Photodiode-Based Optical Imaging for Recording Network Dynamics with Single-Neuron Resolution in Non-Transgenic Invertebrates

The development of transgenic invertebrate preparations in which the activity of specifiable sets of neurons can be recorded and manipulated with light represents a revolutionary advance for studies of the neural basis of behavior. However, a downside of this development is its tendency to focus investigators on a very small number of &#8220;designer&#8221; organisms (e.g., C. elegans and Drosophila), potentially negatively impacting the pursuit of comparative studies across many species, which is needed for identifying general principles of network function. The present article illustrates how optical recording with voltage-sensitive dyes in the brains of non-transgenic gastropod species can be used to rapidly (i.e., within the time course of single experiments) reveal features of the functional organization of their neural networks with single-cell resolution. We outline in detail the dissection, staining, and recording methods used by our laboratory to obtain action potential traces from dozens to ~150 neurons during behaviorally relevant motor programs in the CNS of multiple gastropod species, including one new to neuroscience &#8211; the nudibranch Berghia stephanieae. Imaging is performed with absorbance voltage-sensitive dyes and a 464-element photodiode array that samples at 1,600 frames/second, fast enough to capture all action potentials generated by the recorded neurons. Multiple several-minute recordings can be obtained per preparation with little to no signal bleaching or phototoxicity. The raw optical data collected through the methods described can subsequently be analyzed through a variety of illustrated methods. Our optical recording approach can be readily used to probe network activity in a variety of non-transgenic species, making it well suited for comparative studies of how brains generate behavior.

This protocol presents a method for imaging neuronal population activity with single-cell resolution in non-transgenic invertebrate species using absorbance voltage-sensitive dyes and a photodiode array. This approach enables a rapid workflow, wherein imaging and analysis can be pursued over the course of a single day.

The development of transgenic invertebrate preparations in which the activity of specifiable sets of neurons can be recorded and manipulated with light ...