The authors would like to thank Eleanor Fleming and Jessica Chen for helpful discussions. Funding was provided by National Institute of Allergy and Infectious Diseases (NIH-NIAID) Award R15AI079685 and a Research Corporation Cottrell College Science Award. Additional student support was provided by the Howard Hughes Medical Institute and the Staley fund.

1. Preparation of Large Unilamellar Lipid Vesicles (LUVs)
<ol>
 <li>Prepare LUVs to serve as cell membrane mimics for the assay19.</li>
 <li>Mix phosphatidylcholine (POPC, 760.10 g/mol), phosphatidylglycerol (POPG. 770.99 g/mol), 5-dimethylaminonaphthalene-1-sulfonyl phosphatidylethanolamine (DNS-POPE, 994.350 g/mol) (Avanti Polar Lipids) dissolved in chloroform in a 50:45:5 ratio. As every assay requires the separate preparation of both experimental and control LUVs, two lipid cake vials should be prepared. 0.376 mg of total lipid (0.188 mg POPC, 0.169 mg POPG, and 0.019 mg DNS-POPE) is usually sufficient to make enough vesicles for a single translocation trial.</li>
 <li>Evaporate off chloroform using an N2 gas stream.</li>
 <li>Dry the lipid cake 9-14 hours in a vacuum desiccator.</li>
 <li>Prepare a stock 10 mM HEPES buffer solution (10 mM HEPES, 238.3 g/mol, 45 mM NaCl, 58.44 g/mol, 1 mM EDTA, 372.24 g/mol pH 7.4). HEPES buffer is stored at 4&deg;C.</li>
 <li>Prepare a stock solution of 0.4 mM porcine trypsin (23.3 kg/mol) in 10 mM HEPES buffer prepared above. Trypsin is stored at -20&deg;C and, in our practice, usually does not undergo more than 2 freeze-thaw cycles before use.</li>
 <li>Prepare a stock solution of 4.0 mM Bowman-Birk trypsin inhibitor (8 kg/mol) in 10 mM HEPES buffer solution and store at -20&deg;C. In our use, this stock is replenished monthly and usually does not undergo more than 3 freeze/thaw cycles before it is made again.</li>
 <li>Reconstitute experimental multilamellar vesicles (MLVs) in a 0.2 mM trypsin solution by adding an equal volume of 0.4 mM trypsin stock and 10 mM HEPES buffer solution to the dried lipid cake. For a single sample of 0.376 mg total lipid, a mixture of 150 &mu;L of trypsin solution and 150 &mu;L of HEPES buffer is sufficient.</li>
 <li>Reconstitute control MLVs in a solution with 0.2 mM trypsin and 2.0 mM Bowman-Birk trypsin inhibitor by adding an equal volume of 0.4 mM trypsin stock and 4.0 mM trypsin inhibitor to the dried lipid cakes. For a single sample of 0.376 mg total lipid, a mixture of 150 &mu;L of trypsin solution and 150 &mu;L of trypsin inhibitor is sufficient.</li>
 <li>Transfer MLV solutions from glass vials to 1.7 ml microcentrifuge tube.</li>
 <li>Subject MLV solutions to 5 freeze/thaw cycles in liquid nitrogen.</li>
 <li>Prepare lipid extruder (Avanti Polar Lipids) by placing two filter supports moistened in HEPES buffer on each Teflon extruder piece. Place a nuclepore track etched membrane with 0.1 &mu;m pores (Whatman, United Kingdom) in between the two Teflon extruder pieces, and place the extruder pieces inside the metal extruder canister. Place the donut spacer over the two extruder pieces before tightening the metal top of the extruder canister and placing it in the holder.</li>
 <li>Fill the void volume within the extruder with 500 &mu;L 10 mM HEPES buffer using a 250 &mu;L glass syringe to prevent loss of vesicle sample.</li>
 <li>Load each MLV sample into the extruder and push the sample through the membrane at least 21 times to ensure uniform unilamellar vesicle size. LUVs are obtained from the MLV solution following extrusion.</li>
 <li>Store LUVs at 4&deg;C and use within 48 hours.</li>
</ol>
2. Quantifying LUV Concentration
<ol>
 <li>LUV concentration was quantified by determining the total phosphorus content in the sample20.</li>
 <li>Prepare 8.9 N H2SO4 in nanopure water and store at room temperature.</li>
 <li>Prepare a 2.5% w/v ammonium molybdate VI (588.04 g/mol) solution in nanopure water. Prepare a 10% w/v ascorbic acid (176.1 g/mol) solution in nanopure water. Protect the ascorbic acid solution from light exposure by covering the tube with foil. Store both solutions at 4 &deg;C for up to 2 months.</li>
 <li>Prepare 0.65 mM phosphate buffer solution (monobasic anhydrous sodium phosphate, 120 g/mol).</li>
 <li>Prepare six standard tubes. Standards should contain 0.0 &mu;L, 50 &mu;L (0.0325 &mu;moles), 100 &mu;L (0.065 &mu;moles), 175 &mu;L (0.114 &mu;moles), 250 &mu;L (0.163 &mu;moles), or 350 &mu;L (0.228 &mu;moles) of phosphorus solution.</li>
 <li>Prepare LUV sample tubes in triplicate. Each sample tube should contain approximately 0.1 &mu;moles of LUVs. Experimental and control LUV concentrations must be measured separately.</li>
 <li>To correct for any dried phosphorus salts in the Bowman-Birk inhibitor powder, a control LUV blank must be prepared. The blank should contain the same volume of the 0.2 mM trypsin/2.0 mM Bowman-Birk inhibitor solution as was placed in the control LUV sample tube.</li>
 <li>Place 450 &mu;L of 8.9 N H2SO4 into each of the sample, standard, and blank tubes.</li>
 <li>Heat samples, standards, and blanks for 25 minutes at 175-210 &deg;C in an oven.</li>
 <li>Remove all tubes and allow them to cool. Add 150 &mu;L of 30% H2O2 to each of the tubes.</li>
 <li>Heat samples, standards, and blanks for 30 minutes at 175-210 &deg;C.</li>
 <li>Remove the samples, standards, and blanks from heat. If any of the solutions are not yet colorless, add 50 &mu;L H2O2 to all the tubes and heat for an additional 15 minutes at 175-210 &deg;C.</li>
 <li>Add 3.9 mL nanopure H2O, 0.5 mL ammonium molybdate solution and 0.5 mL ascorbic acid solution to each tube.</li>
 <li>Heat all tubes for 5-7 minutes in a boiling water bath.</li>
 <li>Measure the absorbance of each standard and sample at 820 nm using an Agilent 8453 UV-Visible spectrophotometer. The tube containing 0.0 mmoles phosphorus should be used as the blank for all standards and experimental LUV samples. The tube containing the 0.2 mM trypsin/2.0 mM Bowman-Birk inhibitor solution should be used as the blank for the control LUV sample.</li>
 <li>A linear absorbance vs. phosphorus content standard curve can be produced and used to calculate the total phosphorus content and LUV concentration of the samples.</li>
</ol>
3. Preparing Peptide Solutions
<ol>
 <li>Dissolve peptide in nanopure H2O. Peptides used in this assay as described must contain one tryptophan residue.</li>
 <li>Measure the absorbance of the peptide solution at 282 nm in triplicate. Calculate peptide concentration using the molar extinction coefficient for tryptophan of 5700 M-1cm-1.</li>
 <li>Dilute peptide in nanopure water to a final concentration of 30 &mu;M. Store at -20&deg;C</li>
</ol>
4. Translocation Quantification
<ol>
 <li>Prepare experimental and control samples in 1.7 mL microcentrifuge tubes. Add enough LUVs to each solution for a final concentration of 250 &mu;M. Add enough Bowman-Birk inhibitor solution so that inhibitor is a final concentration 10X greater than the trypsin concentration. Add enough 10 mM HEPES buffer to bring the final solution volume to 450 &mu;L.</li>
 <li>Place 50 &mu;L peptide solution into a Starna micro quartz fluorometer cell. The ratio between LUVs and peptide is high enough to ensure that virtually all intact peptides are in close enough proximity to a dansyl group to give a FRET signal even if none have translocated into the vesicle. This is consistent with the similar FRET observed in control conditions for peptides that do and do not translocate into vesicles.</li>
 <li>Add the experimental or control sample to the peptide solution in the cuvette. This should bring the final peptide concentration to 3 &mu;M.</li>
 <li>Begin a 25 minute fluorescent kinetics experiment immediately upon addition of experimental or control LUV solution, taking a fluorescence reading at least once per second. Set the excitation wavelength at 280 nm, the emission wavelength at 525 nm, and the temperature to 25 &deg;C. Using a Cary Eclipse Fluorescence Spectrophotometer, we set the PMT sensitivity to high.</li>
</ol>
5. Generating a Quantitative Translocation Ratio
<ol>
 <li>Define initial fluorescence (Fo) as the fluorescence reading after fifteen seconds of data collection. In our instrument setup, we have found that the first fifteen seconds of fluorescence data collected are unreliable due to mixing, closing the sample chamber, and other perturbations to the system upon sample addition. Divide each subsequent reading by Fo to obtain relative fluorescence (F/F0) at each time point.</li>
 <li>Average all relative fluorescence readings from the last minute of the experiment to obtain a final average relative fluorescence <img alt="[F/Fo]avg" src="/files/ftp_upload/3571/3571eq1.jpg" />.</li>
 <li>Divide the <img alt="[F/Fo]avg" src="/files/ftp_upload/3571/3571eq1.jpg" /> for control samples by the <img alt="[F/Fo]avg" src="/files/ftp_upload/3571/3571eq1.jpg" /> for experimental samples to obtain a corrected final fluorescence value.</li>
</ol>
6. Representative Results
Figure 2 shows the results of this assay for a representative peptide that showed robust translocation. The signal in this experiment (black trace) shows a marked drop in FRET signal over time. However, it is important to control for the potential loss of FRET signal due to incomplete trypsin inhibition or other factors unrelated to translocation ability. To this end, we always also measure the FRET signal between peptide and LUVs containing both trypsin and Bowman-Birk trypsin inhibitor (gray trace, Figure 2). In our hands, it has been important to perform this control for every peptide every time an experiment is run. This allows us to clearly correct for any changes in signal due to any differences between lipid vesicle preparations or instrument noise, which can be significant for the relatively weak fluorescent signals typically observed at these concentrations.
The importance of the control experiment using LUVs containing trypsin inhibitor is highlighted by the data shown in Figure 3. In this case, the peptide signal decreased a similar amount to that in Figure 2 in the experimental sample (black trace). However, the control sample shows a more rapid decrease for this peptide, so its net translocation is lower.
Section 5 of our protocol describes a straightforward method for quantifying translocation from this experiment. Higher translocation ratios are indicative of peptides that translocate efficiently; the translocation ratio for the cell penetrating peptide shown in Figure 2 is 1.16, while weakly translocating peptides have translocation ratios closer to 1. In our experience, the standard error for three independent experiments performed with different vesicle preparations is in the range of 0.01 to 0.06.
<img alt="Figure 1" src="/files/ftp_upload/3571/3571fig1.jpg" /> 
Figure 1. Schematic of translocation assay. Experimental LUV samples (A) are doped with fluorescent dansyl POPE (black bars) and contain encapsulated trypsin (scissors). Trypsin inhibitor (red circle) is used to inhibit trypsin outside the LUVs. The LUVs are exposed to peptide (purple). Peptide association with LUV membranes yields a FRET signal (green) that decreases as translocting peptides encounter uninhibited internal trypsin. Both trypsin and trypsin inhibitor are encapsulated in control LUV samples (B) to measure decreases in FRET signal unrelated to translocation.
<img alt="Figure 2" src="/files/ftp_upload/3571/3571fig2.jpg" /> 
Figure 2. Representative data for a cell-penetrating peptide. A representative translocating antimicrobial pepide shows a significant drop in FRET signal compared to its control.
<img alt="Figure 3" src="/files/ftp_upload/3571/3571fig3.jpg" /> 
Figure 3. Representative data highlighting the importance of the control. Incomplete inhibition of tryptic digestion of a non-translocating peptide leads to a drop in FRET signal over time in both experimental and control LUV solutions. Because the difference between the control and experimental traces is relatively small, this mutant is characterized as a weakly translocating peptide.

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No conflicts of interest declared.

The protocol presented here can be used to assess the relative change in the concentration of peptides inside and outside of lipid vesicles. These changes are related to translocation ability. This protocol can be used to identify cell-penetrating peptides with potential as drug delivery vectors. As interest in cell-penetrating peptides grows, it will be interesting to see how methods that directly measure the translocation event are developed and used in a quantitative manner.
In our lab, we...

<table border="1">
<tbody>
<tr>
<td>Name of the reagent</td>
<td>Company</td>
<td>Catalogue number</td>
</tr>
<tr>
<td>16:0-18:1 PG</td>
<td>Avanti Polar Lipids</td>
<td>840457C</td>
</tr>
<tr>
<td>1:18 Dansyl PE</td>
<td>Avanti Polar Lipids</td>
<td>810330C</td>
</tr>
<tr>
<td>16:0-18:1 PC</td>
<td>Avanti Polar Lipids</td>
<td>850457C </td>
</tr>
<tr>
<td>Porcine trypsin</td>
<td>Sigma</td>
<td>T-0303</td>
</tr>
<tr>
<td>Bowman-Birk trypsin/chymotrypsin inhibitor</td>
<td>Sigma</td>
<td>T-9777</td>
</tr>
<tr>
<td>Mini-extruder</td>
<td>Avanti Polar Lipids</td>
<td>610000</td>
</tr>
<tr>
<td>Ammonium molybdate (para)</td>
<td>Alfa Aesar</td>
<td>10811</td>
</tr>
<tr>
<td>L-ascorbic acid</td>
<td>Sigma</td>
<td>A1417</td>
</tr>
<tr>
<td>Hydrogen Peroxide, 30% solution</td>
<td>Mallinckrodt chemicals</td>
<td>5240-05</td>
</tr>
<tr>
<td>HEPES</td>
<td>Sigma</td>
<td>H-3375</td>
</tr>
<tr>
<td>NaCl</td>
<td>Sigma</td>
<td>S-9625</td>
</tr>
<tr>
<td>EDTA</td>
<td>Sigma</td>
<td>E5134</td>
</tr>
<tr>
<td>NaH2PO4</td>
<td>Sigma</td>
<td>S-0751</td>
</tr>
</tbody>
</table>

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.

Watch this Scientific Journal Video about Measuring Peptide Translocation into Large Unilamellar Vesicles at JoVE.com

Measuring Peptide Translocation into Large Unilamellar Vesicles

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 ...

measuring-peptide-translocation-into-large-unilamellar-vesicles

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Biology

13.8K Views.  Wellesley College. 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.

Video: Measuring Peptide Translocation into Large Unilamellar Vesicles

Large-Scale Screens of Metagenomic Libraries

Metagenomic libraries archive large fragments of contiguous genomic sequences from microorganisms without requiring prior cultivation. Generating a streamlined procedure for creating and screening metagenomic libraries is therefore useful for efficient high-throughput investigations into the genetic and metabolic properties of uncultured microbial assemblages. Here, key protocols are presented on video, which we propose is the most useful format for accurately describing a long process that alternately depends on robotic instrumentation and (human) manual interventions. First, we employed robotics to spot library clones onto high-density macroarray membranes, each of which can contain duplicate colonies from twenty-four 384-well library plates. Automation is essential for this procedure not only for accuracy and speed, but also due to the miniaturization of scale required to fit the large number of library clones into highly dense spatial arrangements. Once generated, we next demonstrated how the macroarray membranes can be screened for genes of interest using modified versions of standard protocols for probe labeling, membrane hybridization, and signal detection. We complemented the visual demonstration of these procedures with detailed written descriptions of the steps involved and the materials required, all of which are available online alongside the video.

Metagenomic libraries archive large fragments of contiguous genomic sequences from microorganisms without requiring prior cultivation.  Generating a ...

Large Volume (20L+) Filtration of Coastal Seawater Samples

The workflow begins with the collection of coastal marine waters for downstream microbial community, nutrient and trace gas analyses. For this method, samples were collected from the deck of the HMS John Strickland operating in Saanich Inlet. This video documents large volume (&ge;20 L) filtration of microbial biomass, ranging between 0.22&mu;m and 2.7&mu;m in diameter, from the water column. Two 20L samples can be filtered simultaneously using a single pump unit equipped with four rotating heads. Filtration is done in the field on extended trips, or immediately upon return for day trips. It is important to record the amount of water passing through each sterivex filter unit. To prevent biofilm formation between sampling trips, all filtration equipment must be rinsed with dilute HCl and deionized water and autoclaved immediately after use. This procedure will take approximately 5 hours plus an additional hour for clean up.

Large Volume 20L+ Filtration of Coastal Seawater Samples

This video documents large volume (&ge;20 L) filtration of microbial biomass, ranging between 0.22&mu;m and 2.7&mu;m in diameter, from the water column.

The workflow begins with the collection of coastal marine waters for downstream microbial community, nutrient and trace gas analyses. For this method, ...

Large Insert Environmental Genomic Library Production

The vast majority of microbes in nature currently remain inaccessible to traditional cultivation methods. Over the past decade, culture-independent environmental genomic (i.e. metagenomic) approaches have emerged, enabling researchers to bridge this cultivation gap by capturing the genetic content of indigenous microbial communities directly from the environment. To this end, genomic DNA libraries are constructed using standard albeit artful laboratory cloning techniques. Here we describe the construction of a large insert environmental genomic fosmid library with DNA derived from the vertical depth continuum of a seasonally hypoxic fjord. This protocol is directly linked to a series of connected protocols including coastal marine water sampling [1], large volume filtration of microbial biomass [2] and a DNA extraction and purification protocol [3]. At the outset, high quality genomic DNA is end-repaired with the creation of 5 -phosphorylated blunt ends. End-repaired DNA is subjected to pulsed-field gel electrophoresis (PFGE) for size selection and gel extraction is performed to recover DNA fragments between 30 and 60 thousand base pairs (Kb) in length. Size selected DNA is purified away from the PFGE gel matrix and ligated to the phosphatase-treated blunt-end fosmid CopyControl vector pCC1 (EPICENTRE <a href="http://www.epibio.com/item.asp?ID=385">http://www.epibio.com/item.asp?ID=385</a>). Linear concatemers of pCC1 and insert DNA are subsequently headfull packaged into phage particles by lambda terminase, with subsequent infection of phage-resistant E. coli cells. Successfully transduced clones are recovered on LB agar plates under antibiotic selection and archived in 384-well plate format using an automated colony picking robot (Qpix2, GENETIX). The current protocol draws from various sources including the CopyControl Fosmid Library Production Kit from EPICENTRE and the published works of multiple research groups [4-7]. Each step is presented with best practice in mind. Whenever possible we highlight subtleties in execution to improve overall quality and efficiency of library production. The whole process of fosmid library production and automated colony picking takes at least 7-10 days as there are many incubation steps included. However, there are several stopping points possible which are mentioned within the protocol.

Construction of a fosmid library with environmental genomic DNA isolated from the vertical depth continuum of a seasonally hypoxic fjord is described. The resulting clone library is picked into 384-well plates and archived for downstream sequencing and functional screening by the application of an automated colony picking system.

The vast majority of microbes in nature currently remain inaccessible to traditional cultivation methods. Over the past decade, culture-independent ...

Mutagenesis and Functional Analysis of Ion Channels Heterologously Expressed in Mammalian Cells

We will demonstrate how to study the functional effects of introducing a point mutation in an ion channel. We study G protein-gated inwardly rectifying potassium (referred to as GIRK) channels, which are important for regulating the excitability of neurons. There are four different mammalian GIRK channel subunits (GIRK1-GIRK4) - we focus on GIRK2 because it forms a homotetramer. Stimulation of different types of G protein-coupled receptors (GPCRs), such as the muscarinic receptor (M2R), leads to activation of GIRK channels. Alcohol also directly activates GIRK channels. We will show how to mutate one amino acid by specifically changing one or more nucleotides in the cDNA for the GIRK channel. This mutated cDNA sequence will be amplified in bacteria, purified, and the presence of the point mutation will be confirmed by DNA sequencing. The cDNAs for the mutated and wild-type GIRK channels will be transfected into human embryonic kidney HEK293T cells cultured in vitro. Lastly, whole-cell patch-clamp electrophysiology will be used to study the macroscopic potassium currents through the ectopically expressed wild-type or mutated GIRK channels. In this experiment, we will examine the effect of a L257W mutation in GIRK2 channels on M2R-dependent and alcohol-dependent activation.

We will demonstrate how to study the effect of a single point mutation on the function of an ion channel.

We will demonstrate how to study the functional effects of introducing a point mutation in an ion channel. We study G protein-gated inwardly rectifying ...

Quantitative Measurement of GLUT4 Translocation to the Plasma Membrane by Flow Cytometry

Glucose is the main source of energy for the body, requiring constant regulation of its blood concentration. Insulin release by the pancreas induces glucose uptake by insulin-sensitive tissues, most notably the brain, skeletal muscle, and adipocytes. Patients suffering from type-2 diabetes and/or obesity often develop insulin resistance and are unable to control their glucose homeostasis. New insights into the mechanisms of insulin resistance may provide new treatment strategies for type-2 diabetes.
The GLUT family of glucose transporters consists of thirteen members distributed on different tissues throughout the body1. Glucose transporter type 4 (GLUT4) is the major transporter that mediates glucose uptake by insulin sensitive tissues, such as the skeletal muscle. Upon binding of insulin to its receptor, vesicles containing GLUT4 translocate from the cytoplasm to the plasma membrane, inducing glucose uptake. Reduced GLUT4 translocation is one of the causes of insulin resistance in type-2 diabetes2,3.
The translocation of GLUT4 from the cytoplasm to the plasma membrane can be visualized by immunocytochemistry, using fluorophore-conjugated GLUT4-specific antibodies.
Here, we describe a technique to quantify total amounts of GLUT4 translocation to the plasma membrane of cells during a chosen duration, using flow cytometry. This protocol is rapid (less than 4 hours, including incubation with insulin) and allows the analysis of as few as 3,000 cells or as many as 1 million cells per condition in a single experiment. It relies on anti-GLUT4 antibodies directed to an external epitope of the transporter that bind to it as soon as it is exposed to the extracellular medium after translocation to the plasma membrane.

This protocol describes a rapid technique to quantify the translocation of GLUT4 from the cytoplasm to the plasma membrane of cells by flow cytometry.

Glucose is the main source of energy for the body, requiring constant regulation of its blood concentration. Insulin release by the pancreas induces ...

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.

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 ...

Infinium Assay for Large-scale SNP Genotyping Applications

Genotyping variants in the human genome has proven to be an efficient method to identify genetic associations with phenotypes. The distribution of variants within families or populations can facilitate identification of the genetic factors of disease. Illumina&#39;s panel of genotyping BeadChips allows investigators to genotype thousands or millions of single nucleotide polymorphisms (SNPs) or to analyze other genomic variants, such as copy number, across a large number of DNA samples. These SNPs can be spread throughout the genome or targeted in specific regions in order to maximize potential discovery. The Infinium assay has been optimized to yield high-quality, accurate results quickly. With proper setup, a single technician can process from a few hundred to over a thousand DNA samples per week, depending on the type of array. This assay guides users through every step, starting with genomic DNA and ending with the scanning of the array. Using propriety reagents, samples are amplified, fragmented, precipitated, resuspended, hybridized to the chip, extended by a single base, stained, and scanned on either an iScan or Hi Scan high-resolution optical imaging system. One overnight step is required to amplify the DNA. The DNA is denatured and isothermally amplified by whole-genome amplification; therefore, no PCR is required. Samples are hybridized to the arrays during a second overnight step. By the third day, the samples are ready to be scanned and analyzed. Amplified DNA may be stockpiled in large quantities, allowing bead arrays to be processed every day of the week, thereby maximizing throughput.

A protocol is described that uses Illumina&#39;s Infinium assays to perform large-scale genotyping. These assays can reliably genotype millions of SNPs across hundreds of individual DNA samples in three days. Once generated, these genotypes can be used to check for associations with a variety of different diseases or phenotypes.

Genotyping variants in the human genome has proven to be an efficient method to identify genetic associations with phenotypes. The distribution of ...

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 ...