We thank Michael J. Bruno, Radda Rusinova and Jon T. Sack for many stimulating discussions. Financial support from NIH, R01GM021342 and ARRA supplement R01GM021342-35S1, and the Josiah Macy, Jr. Foundation to OSA; the Tri-I CMB program for HII; and The Iris L. and Leverett S. Woodworth Medical Scientist Fellowship and NIH MSTP grant GM07739 for RK.

1. Generate ANTS-filled Liposomes
<ol>
 <li>On day 1, remove organic solvent from lipids.</li> 
	<li>Remove lipid from freezer and let it equilibrate to room temperature.</li> 
	<li>Add 0.6 mL of 25 mg/mL (1,2-dierucoyl-sn-glycero-3-phosphocholine) lipid in chloroform solution to a 25 mL round bottom flask.</li> 
	<li>Continually rotate the flask while drying under nitrogen, until all the chloroform has evaporated and a thin white film of lipid coats the entire lower half of the flask.</li> 
	<li>Dry in a dessicator under vacuum overnight.</li>
</ol>
<ol>
 <li>On day 2, prepare the sample with the fluorophore that will be incorporated into the vesicles.</li> 
	<li>Rehydrate in 100 mM NaNO3, 25 mM ANTS (Na salt), 10 mM HEPES. Use HNO3 or NaOH to adjust pH to 7.0. Each ANTS molecule has 2 Na+, for a total of 150 mM [Na+]. Add 1.671 mL of electrolyte to get a 10 mM lipid suspension.</li> 
	<li>Parafilm and vortex suspension thoroughly.</li> 
	<li>Protect sample from light with foil and let it age at room temperature overnight.</li>
</ol>
<ol>
 <li>On Day 3, make large unilamellar vesicles and remove all the fluorophore from outside the vesicles.</li> 
	<li>Sonicate for 1 min in a low-power sonicator.</li> 
	<li>Freeze-thaw sample 5-6 times, each time with 5 minutes on dry ice and 5 minutes in warm (~50 &deg;C) water.</li> 
	<li>Extrude the lipid suspension using an Avanti mini-extruder. Set up the mini-extruder with a 0.1 &mu;m polycarbonate filter and filter supports (see extruder manual for details). Extrude the suspension back and forth 21 times such that the suspension ends up on the opposite syringe. When multiple batches are extruded, always remember to load samples with the same syringe. During extrusion the cloudy initial lipid suspension should become almost translucent (it will still be yellow due to the ANTS fluorophore). If the extrusion suddenly becomes easier and less pressure is needed to move through filter, the filter may have ruptured and will need to be replaced.</li> 
	<li>Remove external ANTS by running the extruded suspension over a PD-10 desalting column using a gravity protocol. Equilibrate column with 20-30 mL of Na-buffer (140 mM NaNO3, 10 mM HEPES, pH 7.0, remember to use HNO3 or NaOH to adjust pH). Add 1.5 mL of the extruded lipid suspension to the column and let it seep in. Bring the total volume of sample to 2.5 mL by adding 1 mL Na-buffer. After the buffer is fully incorporated into the column and nothing is leaking out, elute the liposomes with 3 mL of Na-buffer and collect the resulting sample. The resulting ANTS filled LUV stock solution should contain about 4-5 mM lipids and appear translucent milky white. Protect the sample from light with foil.</li> 
	<li>If the stock solution is not used immediately, store at 13 &deg;C for a maximum of 7 days. Warning: do not freeze or cool the lipid solution below its liquid- to crystal-phase transition temperature (~12 &deg;C), as this will destroy the vesicles' integrity and the fluorophore will leak out.</li>
</ol>
2. Mix Fluorescence Solution
<ol>
 <li>24 hours prior to use, dilute liposomes and incubate with gramicidin (at 13 &deg;C). Equilibration of gramicidin between the lipid vesicles' inner and outer monolayers is a slow process, requiring a long incubation.</li> 
	<li>Thoroughly vortex the ANTS-filled LUV stock solution as the LUVs have a density >1 g/mL and will therefore sediment in the solutions.</li> 
	<li>Mix the ANTS-filled LUV stock 1:20 in Na-buffer. Mix all liposomes for use in one day of experiments together in a single flask to enhance sample uniformity.</li> 
	<li>Separate out 3/4 of the liposomes and add 260 nM gramicidin pre-diluted in DMSO (500 &mu;g/mL). </li> 
	<li>To the remaining 1/4 add the same volume of DMSO (without gramicidin) to keep the solvent concentration constant in all samples.</li>
</ol>
3. Setting up the Fluorescence Instrument
<ol>
 <li>We use an Applied Photophysics SX.20 stopped-flow spectrofluorometer with temperature control to measure the rate of fluorescence quenching.</li> 
	<li>Turn the instruments on an hour ahead of time to allow for warm up. The components are: computer, electronic control unit, water-bath (both cooler and heater), lamp power supply and nitrogen tank.</li> 
	<li>Set up the recording conditions using the Pro-Data SX software.</li> 
	<li>Set the monochromator excitation wavelength to 352 nm and slit width apertures for excitation to 1/1.</li> 
	<li>Use a &lambda;=455 nm high-pass filter to record the emission.</li> 
	<li>Use the "Pressure Hold" setting, which enables the instrument to maintain the nitrogen pressure during the recording and thus prevents pressure oscillations (and recording artifacts) during the first few milliseconds.</li> 
	<li>Set "Time" to 1 s and "Points" to 5000.</li> 
	<li>Use the repeat box and adjust the number of repeats, normally we use 9 and 13 repeats for buffer and quencher runs, respectively.</li> 
	<li>Water-bath temperature should be set to 25&deg;C; the temperature should be monitored in the SX software.</li> 
	<li>Set the drive volume to 120 &mu;L, meaning that each time the instrument runs a sample, it mixes 60 &mu;L from each of the two syringes. With this instrumental setup, a 120 &mu;L volume gives a dead time of &asymp; 1.2 ms.</li> 
	<li>Adjust the high voltage (gain) on the fluorescence detector for the ANTS-liposome's output to be around 8. For a standard experimental mixture the gain should be 400-420 V. Some drugs are fluorescent, such that they can saturate the fluorescence signal. In these cases, gain/voltage should be decreased.</li> 
	<li>Always load the fluorescent sample into the left syringe and buffer/quencher in the right syringe. Make sure to thoroughly vortex all samples containing liposomes before loading, and be aware that the LUVs with time will settle in the syringe.</li>
</ol>
4. Doing an Experiment
<ol>
 <li>Prepare a sample of ANTS-loaded LUVs with either solvent or compound, and load into the spectrofluorometer together with either Na-buffer or Tl-quencher (50 mM TlNO3, 94 mM NaNO3, 10 mM HEPES at pH 7.0).</li> 
	<li>Use 1.5 mL diluted ANTS-LUV solution, with or without gramicidin, prepared the previous day.</li> 
	<li>Add the compound at the desired concentration or the solvent for controls. Keep the solvent concentration to a minimum and constant across all the samples. The concentrations will need to be adjusted with any new (unknown) compound to achieve the desired dose-response range. Incubate for 10 min at 25&deg;C in the dark.</li> 
	<li>Vortex the LUV sample and load into the left syringe. Load the Na-buffer or Tl-quencher into the right syringe.</li> 
	<li>Thoroughly remove air bubbles from both samples by pushing the syringes back and forth.</li> 
	<li>Make sure both syringes are loaded equally before closing the valves. Unequal loading will result in pre-mixing of solutions before they reach the recording chamber.</li> 
	<li>For the very first sample, record the fluorescence with Na-buffer, repeat 4 times, adjust the high voltage (gain) as necessary, and repeat 5 times.</li> 
	<li>For the remaining samples, record 9 repeats with Na-buffer.</li> 
	<li>Replace the Na-buffer with the Tl-quencher and record 13 repeats.</li> 
	<li>Rinse with water and continue with next sample. As controls we use samples with: no gramicidin and with solvent, no gramicidin and with the maximum compound concentration, with both gramicidin and solvent.</li>
</ol>
5. Analyzing Data
<ol>
 <li>Transfer the results to the analysis computer. Read all the data into MATLAB for analyzing. For each sample, read in all the buffer and quencher repeats, excluding the first four in each case, as those contain mixing artifacts. Assuming a drive volume of 120 &mu;L it takes a little less than four shots to completely clear out the previous mixture from the stopped-flow tubing.</li> 
	<li>The buffer repeats for all the samples should be very similar. If there is a significant shift in the fluorescence signal, which depends on the compound concentration, then the compound itself may be fluorescent and it may be necessary to first subtract this extra fluorescence signal before normalizing the samples.</li> 
	<li>Manually go through the traces and remove "bad" repeats: repeats containing spikes or deviations from multi-exponentiality due to mixing artifacts and/or bubbles.</li> 
	<li>Normalize the quencher repeats to the average starting value of the buffer repeats for each sample. Combine averages of all samples into one graph for visualization.</li> 
	<li>Traces recorded with no gramicidin should all be similar and show almost no fluorescence quenching; there may be a slow reduction in the fluorescence signal due to slow leakage of Tl+ through the vesicles bilayer 8. If the sample with no gramicidin and no modifier shows significant quenching, then the vesicles have deteriorated and should not be used further. If the samples with no gramicidin but with the modifier show significant quenching, then the modifier perturbs the vesicles bilayer to such an extent that it allows the fluorophore or quencher to cross the bilayer.</li> 
	<li>If the compound alters lipid bilayer properties, as sensed by gramicidin channels, the fluorescence time course will be visibly altered.</li> 
	<li>For each sample, a stretched exponential <img src="/files/ftp_upload/2131/2131eg1.jpg" alt="Equation 1" /> e.g. 4) is fit to the first 2-100 ms of the normalized fluorescence quenching curve of the individual repeats and the rate <img src="/files/ftp_upload/2131/2131eg2.jpg" alt="Figure 2" />4 calculated at 2 ms. Averages and standard deviations are calculated for a given sample from all the individual repeats.</li> 
	<li>Finally, determine the relative change in quench rate by normalizing to the closest control sample in time that has gramicidin and no modifier.</li>
</ol>
6. Representative Results
<img src="/files/ftp_upload/2131/2131fig1.jpg" alt="Figure 1" /> Figure 1: The essentials of the fluorescence quench-based assay to detect changes in lipid bilayer properties. Top left: A zoom-in on a single lipid vesicle with ANTS plus NaNO3 on the inside and NaNO3 plus TlNO3 on the outside. Top right: The fluorescence signal recorded using (form top to bottom) ANTS-filled vesicles without quencher, with quencher, with quencher and pre-doped with 87, 260 and 780 nM gramicidin. Bottom: Schematic representation of the stopped-flow mixing chamber.
<img src="/files/ftp_upload/2131/2131fig2.jpg" alt="Figure 2" /> Figure 2: A screen shot from the Pro-Data SX software illustrating the various panels referenced in the description of the experimental setup.
<img src="/files/ftp_upload/2131/2131fig3.jpg" alt="Figure 3" /> Figure 3: Multiple repeat determinations of the fluorescence signal from ANTS-loaded LUVs with Na-buffer. The first four repeats are always excluded, as they contain mixing artifacts. The tubing connecting the sample syringes to the mixing cell have a defined volume, therefore the first few repeats will give us a reading of what was previously in the tubing: water for repeat 1 and 2, some combination of water and sample for repeat 3, mostly sample for repeat 4, and just sample for the remaining repeats. 
<img src="/files/ftp_upload/2131/2131fig4.jpg" alt="Figure 4" /> Figure 4: Multiple repeat determinations of the fluorescence signal from ANTS-loaded LUVs with Na-buffer and with Tl-quencher. The first four repeats have been removed from both conditions. Additionally, for the Tl-quencher measurements, repeat 8 needs to be removed due to artifacts, most likely air bubbles.
<img src="/files/ftp_upload/2131/2131fig5.jpg" alt="Figure 5" /> Figure 5: Effect of capsaicin (Cap) on the time course of ANTS fluorescence quenching. (A) Normalized fluorescence signal over 1 s, gray dots denote results from all repeats (n &gt; 5 per condition); red lines denote the average of all repeats. (B) The first 100 ms, gray dots denote results from a single repeat for each condition; red lines are stretched exponential fits (2 - 100 ms) to those repeats. The stippled blue line denotes the 2 ms mark, the time at which the rate of quenching is determined. In both A and B the top trace shows results in the absence of Tl+; the next two traces show results in the absence of gA, with Tl+ &plusmn; Cap; the four lower traces show results with 260 nM gA and Tl+, where the numbers denote [Cap] in &mu;M. The rates of quenching for 0, 10, 30 and 90 &mu;M Cap, as determined by the rate of a stretched exponential, are 36&plusmn;6, 69&plusmn;6, 85&plusmn;8 and 247&plusmn;27 (mean &plusmn; s.d., n &gt; 8), respectively.

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A provisional patent application that covers the methods described in the manuscript has been filed. The co-inventors are HII and OSA.

We have demonstrated a fast fluorescence-based assay for determining the bilayer modifying potential of drugs and other small amphiphiles. Compounds that modify bilayer properties are likely to alter membrane protein function in an indirect, nonspecific manner, possibly contributing to "off-target" drug effects. The assay exploits the power of gramicidin channels as probes for changes in bilayer properties 12 that are sensed by bilayer-spanning proteins. The results obtained using the fluorescence-based ass...

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		<th>Material Name</th>
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		<th>Catalogue Number</th>
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<td>ANTS</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>A-350</td>
<td>&nbsp;</td>
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<td>gramicidin</td>
<td>&nbsp;</td>
<td>Sigma Chemical Co</td>
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<td>1,2-dierucoyl-sn-glycero-3-phosphocholine</td>
<td>&nbsp;</td>
<td>Avanti Polar Lipids</td>
<td>850398C</td>
<td>&nbsp;</td>
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<td>Mini-Extruder kit</td>
<td>&nbsp;</td>
<td>Avanti Polar Lipids</td>
<td>610000</td>
<td>&nbsp;</td>
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gramicidin-based fluorescence assay; for determining small molecules potential for modifying lipid bilayer properties

Many drugs and other small molecules used to modulate biological function are amphiphiles that adsorb at the bilayer/solution interface and thereby alter lipid bilayer properties. This is important because membrane proteins are energetically coupled to their host bilayer by hydrophobic interactions. Changes in bilayer properties thus alter membrane protein function, which provides an indirect way for amphiphiles to modulate protein function and a possible mechanism for "off-target" drug effects. We have previously developed an electrophysiological assay for detecting changes in lipid bilayer properties using linear gramicidin channels as probes 3,12. Gramicidin channels are mini-proteins formed by the transbilayer dimerization of two non-conducting subunits. They are sensitive to changes in their membrane environment, which makes them powerful probes for monitoring changes in lipid bilayer properties as sensed by bilayer spanning proteins. We now demonstrate a fluorescence assay for detecting changes in bilayer properties using the same channels as probes. The assay is based on measuring the time-course of fluorescence quenching from fluorophore-loaded large unilamellar vesicles due to the entry of a quencher through the gramicidin channels. We use the fluorescence indicator/quencher pair 8-aminonaphthalene-1,3,6-trisulfonate (ANTS)/Tl+ that has been successfully used in other fluorescence quenching assays 5,13. Tl+ permeates the lipid bilayer slowly 8 but passes readily through conducting gramicidin channels 1,14. The method is scalable and suitable for both mechanistic studies and high-throughput screening of small molecules for bilayer-perturbing, and potential "off-target", effects. We find that results using this method are in good agreement with previous electrophysiological results 12.

Watch this Scientific Journal Video about Gramicidin-based Fluorescence Assay; for Determining Small Molecules Potential for Modifying Lipid Bilayer Properties at JoVE.com

Gramicidin-based Fluorescence Assay; for Determining Small Molecules Potential for Modifying Lipid Bilayer Properties

We introduce a fast fluorescence-based assay that monitors the rate of fluorescence quenching as a measure of gramicidin channel activity. The gramicidin channels are used as molecular force transducers to monitor changes in lipid bilayer properties as sensed by bilayer spanning proteins.

Many drugs and other small molecules used to modulate biological function are amphiphiles that adsorb at the bilayer/solution interface and thereby alter ...

gramicidin-based-fluorescence-assay-for-determining-small-molecules

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12.7K Views.  Weill Cornell Medical College. We introduce a fast fluorescence-based assay that monitors the rate of fluorescence quenching as a measure of gramicidin channel activity. The gramicidin channels are used as molecular force transducers to monitor changes in lipid bilayer properties as sensed by bilayer spanning proteins.

Video: Gramicidin-based Fluorescence Assay; for Determining Small Molecules Potential for Modifying Lipid Bilayer Properties

Application of a C. elegans Dopamine Neuron Degeneration Assay for the Validation of Potential Parkinson's Disease Genes

Improvements to the diagnosis and treatment of Parkinson's disease (PD) are dependent upon knowledge about susceptibility factors that render populations at risk. In the process of attempting to identify novel genetic factors associated with PD, scientists have generated many lists of candidate genes, polymorphisms, and proteins that represent important advances, but these leads remain mechanistically undefined. Our work is aimed toward significantly narrowing such lists by exploiting the advantages of a simple animal model system. While humans have billions of neurons, the microscopic roundworm Caenorhabditis elegans has precisely 302, of which only eight produce dopamine (DA) in hemaphrodites. Expression of a human gene encoding the PD-associated protein, alpha-synuclein, in C. elegans DA neurons results in dosage and age-dependent neurodegeneration.
Worms expressing human alpha-synuclein in DA neurons are isogenic and express both GFP and human alpha-synuclein under the DA transporter promoter (Pdat-1). The presence of GFP serves as a readily visualized marker for following DA neurodegeneration in these animals. We initially demonstrated that alpha-synuclein-induced DA neurodegeneration could be rescued in these animals by torsinA, a protein with molecular chaperone activity 1. Further, candidate PD-related genes identified in our lab via large-scale RNAi screening efforts using an alpha-synuclein misfolding assay were then over-expressed in C. elegans DA neurons. We determined that five of seven genes tested represented significant candidate modulators of PD as they rescued alpha-synuclein-induced DA neurodegeneration 2. Additionally, the Lindquist Lab (this issue of JoVE) has performed yeast screens whereby alpha-synuclein-dependent toxicity is used as a readout for genes that can enhance or suppress cytotoxicity. We subsequently examined the yeast candidate genes in our C. elegans alpha-synuclein-induced neurodegeneration assay and successfully validated many of these targets 3, 4.
Our methodology involves generation of a C. elegans DA neuron-specific expression vector using recombinational cloning of candidate gene cDNAs under control of the Pdat-1 promoter. These plasmids are then microinjected in wild-type (N2) worms, along with a selectable marker for successful transformation. Multiple stable transgenic lines producing the candidate protein in DA neurons are obtained and then independently crossed into the alpha-synuclein degenerative strain and assessed for neurodegeneration, at both the animal and individual neuron level, over the course of aging.

This video demonstrates how to use C. elegans to assess dopaminergic neuron neurodegeneration as a model for Parkinson's disease. Furthermore, genetic screens are used to identify factors that either enhance degeneration or are neuroprotective.

Improvements to the diagnosis and treatment of Parkinson's disease (PD) are dependent upon knowledge about susceptibility factors that render populations ...

Single Molecule Methods for Monitoring Changes in Bilayer Elastic Properties

Membrane protein function is regulated by the cell membrane lipid composition.  This regulation is due to a combination of specific lipid-protein interactions and more general lipid bilayer-protein interactions. These interactions are particularly important in pharmacological research, as many current pharmaceuticals on the market can alter the lipid bilayer material properties, which can lead to altered membrane protein function.   The formation of gramicidin channels are dependent on conformational changes in gramicidin subunits which are in turn dependent on the properties of the lipid.  Hence the gramicidin channel  current is a reporter of altered properties of the bilayer due to certain compounds.  

Membrane protein function is regulated by the cell membrane lipid composition. This video-article details how to form a patch using bilayer patch electrodes, as well as how to use gramicidin channels as reporters of altered membrane properties.

Membrane protein function is regulated by the cell membrane lipid composition.  This regulation is due to a combination of specific lipid-protein ...

A Fluorescence Microscopy Assay for Monitoring Mitophagy in the Yeast Saccharomyces cerevisiae

Autophagy is important for turnover of cellular components under a range of different conditions. It serves an essential homeostatic function as well as a quality control mechanism that can target and selectively degrade cellular material including organelles1-4. For example, damaged or redundant mitochondria (Fig. 1), not disposed of by autophagy, can represent a threat to cellular homeostasis and cell survival. In the yeast, Saccharomyces cerevisiae, nutrient deprivation (e.g., nitrogen starvation) or damage can promote selective turnover of mitochondria by autophagy in a process termed mitophagy 5-9.
We describe a simple fluorescence microscopy approach to assess autophagy. For clarity we restrict our description here to show how the approach can be used to monitor mitophagy in yeast cells. The assay makes use of a fluorescent reporter, Rosella, which is a dual-emission biosensor comprising a relatively pH-stable red fluorescent protein linked to a pH-sensitive green fluorescent protein. The operation of this reporter relies on differences in pH between the vacuole (pH ~ 5.0-5.5) and mitochondria (pH ~ 8.2) in living cells. Under growing conditions, wild type cells exhibit both red and green fluorescence distributed in a manner characteristic of the mitochondria. Fluorescence emission is not associated with the vacuole. When subjected to nitrogen starvation, a condition which induces mitophagy, in addition to red and green fluorescence labeling the mitochondria, cells exhibit the accumulation of red, but not green fluorescence, in the acidic vacuolar lumen representing the delivery of mitochondria to the vacuole. Scoring cells with red, but not green fluorescent vacuoles can be used as a measure of mitophagic activity in cells5,10-12.

A Fluorescence Microscopy Assay for Monitoring Mitophagy in the Yeast Saccharomyces cerevisiae

A robust approach to monitor the delivery of organelles to the acidic lumen of the yeast vacuole for degradation and recycling is described. The method relies on the specific labeling of target organelles with a genetically encoded dual-emission fluorescence pH-biosensor, and visualization of individual cells using fluorescence microscopy.

Autophagy is important for turnover of cellular components under a range of different conditions. It serves an essential homeostatic function as well as a ...

Substrate Generation for Endonucleases of CRISPR/Cas Systems

The interaction of viruses and their prokaryotic hosts shaped the evolution of bacterial and archaeal life. Prokaryotes developed several strategies to evade viral attacks that include restriction modification, abortive infection and CRISPR/Cas systems. These adaptive immune systems found in many Bacteria and most Archaea consist of clustered regularly interspaced short palindromic repeat (CRISPR) sequences and a number of CRISPR associated (Cas) genes (Fig. 1) 1-3. Different sets of Cas proteins and repeats define at least three major divergent types of CRISPR/Cas systems 4. The universal proteins Cas1 and Cas2 are proposed to be involved in the uptake of viral DNA that will generate a new spacer element between two repeats at the 5' terminus of an extending CRISPR cluster 5. The entire cluster is transcribed into a precursor-crRNA containing all spacer and repeat sequences and is subsequently processed by an enzyme of the diverse Cas6 family into smaller crRNAs 6-8. These crRNAs consist of the spacer sequence flanked by a 5' terminal (8 nucleotides) and a 3' terminal tag derived from the repeat sequence 9. A repeated infection of the virus can now be blocked as the new crRNA will be directed by a Cas protein complex (Cascade) to the viral DNA and identify it as such via base complementarity10. Finally, for CRISPR/Cas type 1 systems, the nuclease Cas3 will destroy the detected invader DNA 11,12 .
These processes define CRISPR/Cas as an adaptive immune system of prokaryotes and opened a fascinating research field for the study of the involved Cas proteins. The function of many Cas proteins is still elusive and the causes for the apparent diversity of the CRISPR/Cas systems remain to be illuminated. Potential activities of most Cas proteins were predicted via detailed computational analyses. A major fraction of Cas proteins are either shown or proposed to function as endonucleases 4.
Here, we present methods to generate crRNAs and precursor-cRNAs for the study of Cas endoribonucleases. Different endonuclease assays require either short repeat sequences that can directly be synthesized as RNA oligonucleotides or longer crRNA and pre-crRNA sequences that are generated via in vitro T7 RNA polymerase run-off transcription. This methodology allows the incorporation of radioactive nucleotides for the generation of internally labeled endonuclease substrates and the creation of synthetic or mutant crRNAs. Cas6 endonuclease activity is utilized to mature pre-crRNAs into crRNAs with 5'-hydroxyl and a 2',3'-cyclic phosphate termini.

CRISPR/Cas systems mediate adaptive immunity in Bacteria and Archaea. Many Cas proteins are proposed to act as endoribonucleases acting on crRNA precursors of varying length. Here we illustrate three different approaches to generate pre-crRNA substrates for the biochemical analysis of Cas endonuclease activity.

The  interaction of viruses and their prokaryotic hosts shaped the evolution of  bacterial and archaeal life. Prokaryotes developed several strategies to ...

Visualization and Analysis of mRNA Molecules Using Fluorescence In Situ Hybridization in Saccharomyces cerevisiae

The Fluorescence in situ Hybridization (FISH) method allows one to detect nucleic acids in the native cellular environment. Here we provide a protocol for using FISH to quantify the number of mRNAs in single yeast cells. Cells can be grown in any condition of interest and then fixed and made permeable. Subsequently, multiple single-stranded deoxyoligonucleotides conjugated to fluorescent dyes are used to label and visualize mRNAs. Diffraction-limited fluorescence from single mRNA molecules is quantified using a spot-detection algorithm to identify and count the number of mRNAs per cell. While the more standard quantification methods of northern blots, RT-PCR and gene expression microarrays provide information on average mRNAs in the bulk population, FISH facilitates both the counting and localization of these mRNAs in single cells at single-molecule resolution.

Visualization and Analysis of mRNA Molecules Using Fluorescence In Situ Hybridization in Saccharomyces cerevisiae

This protocol describes an experimental procedure for performing Fluorescence in situ Hybridization (FISH) for counting mRNAs in single cells at single-molecule resolution.

The Fluorescence in situ Hybridization (FISH) method allows one to detect nucleic acids in the native cellular environment. Here we provide a protocol for ...

Highly Efficient Ligation of Small RNA Molecules for MicroRNA Quantitation by High-Throughput Sequencing

MiRNA cloning and high-throughput sequencing, termed miR-Seq, stands alone as a transcriptome-wide approach to quantify miRNAs with single nucleotide resolution. This technique captures miRNAs by attaching 3&#8217; and 5&#8217; oligonucleotide adapters to miRNA molecules and allows de novo miRNA discovery. Coupling with powerful next-generation sequencing platforms, miR-Seq has been instrumental in the study of miRNA biology. However, significant biases introduced by oligonucleotide ligation steps have prevented miR-Seq from being employed as an accurate quantitation tool. Previous studies demonstrate that biases in current miR-Seq methods often lead to inaccurate miRNA quantification with errors up to 1,000-fold for some miRNAs1,2. To resolve these biases imparted by RNA ligation, we have developed a small RNA ligation method that results in ligation efficiencies of over 95% for both 3&#8217; and 5&#8242; ligation steps. Benchmarking this improved library construction method using equimolar or differentially mixed synthetic miRNAs, consistently yields reads numbers with less than two-fold deviation from the expected value. Furthermore, this high-efficiency miR-Seq method permits accurate genome-wide miRNA profiling from in vivo total RNA samples2.

MicroRNAs (miRNAs) are a widely conserved class of regulatory molecules. Here we describe a miRNA cloning method that relies upon two potent ligation steps followed by high-throughput sequencing. Our method permits accurate genome-wide quantitation of miRNAs.

MiRNA cloning and high-throughput sequencing, termed miR-Seq, stands alone as a transcriptome-wide approach to quantify miRNAs with single nucleotide ...

Determining Membrane Protein Topology Using Fluorescence Protease Protection (FPP)

The correct topology and orientation of integral membrane proteins are essential for their proper function, yet such information has not been established for many membrane proteins. A simple technique called fluorescence protease protection (FPP) is presented, which permits the determination of membrane protein topology in living cells. This technique has numerous advantages over other methods for determining protein topology, in that it does not require the availability of multiple antibodies against various domains of the membrane protein, does not require large amounts of protein, and can be performed on living cells. The FPP method employs the spatially confined actions of proteases on the degradation of green fluorescent protein (GFP) tagged membrane proteins to determine their membrane topology and orientation. This simple approach is applicable to a wide variety of cell types, and can be used to determine membrane protein orientation in various subcellular organelles such as the mitochondria, Golgi, endoplasmic reticulum and components of the endosomal/recycling system. Membrane proteins, tagged on either the N-termini or C-termini with a GFP fusion, are expressed in a cell of interest, which is subject to selective permeabilization using the detergent digitonin. Digitonin has the ability to permeabilize the plasma membrane, while leaving intracellular organelles intact. GFP moieties exposed to the cytosol can be selectively degraded through the application of protease, whereas GFP moieties present in the lumen of organelles are protected from the protease and remain intact. The FPP assay is straightforward, and results can be obtained rapidly.

Determining Membrane Protein Topology Using Fluorescence Protease Protection FPP

Here, we present a protocol to determine the orientation and topology of integral membrane proteins in living cells. This simple protocol relies on selective protease sensitivity of chimeras between the protein of interest and GFP.

The correct topology and orientation of integral membrane proteins are essential for their proper function, yet such information has not been established ...

Luciferase Complementation Imaging Assay in Nicotiana benthamiana Leaves for Transiently Determining Protein-protein Interaction Dynamics

Protein-protein interactions are fundamental mechanisms for relaying signal transduction in most cellular processes; therefore, identification of novel protein-protein interaction pairs and monitoring protein interaction dynamics are of particular interest for revealing how plants respond to environmental factors and/or developmental signals. A plethora of approaches have been developed to examine protein-protein interactions, either in vitro or in vivo. Among them, the recently established luciferase complementation imaging (LCI) assay is the simplest and fastest method for demonstrating in vivo protein-protein interactions. In this assay, protein A or protein B is fused with the amino-terminal or carboxyl-terminal half of luciferase, respectively. When protein A interacts with protein B, the two halves of luciferase will be reconstituted to form a functional and active luciferase enzyme. Luciferase activity can be recorded with a luminometer or CCD-camera. Compared with other approaches, the LCI assay shows protein-protein interactions both qualitatively and quantitatively. Agrobacterium infiltration in Nicotiana benthamiana leaves is a widely used system for transient protein expression. With the combination of LCI and transient expression, these approaches show that the physical interaction between COP1 and SPA1 was gradually reduced after jasmonate treatment.

Luciferase Complementation Imaging Assay in Nicotiana benthamiana Leaves for Transiently Determining Protein-protein Interaction Dynamics

This manuscript describes an easy and rapid experimental procedure for determining protein-protein interactions based on the measurement of luciferase activity.

Protein-protein interactions are fundamental mechanisms for relaying signal transduction in most cellular processes; therefore, identification of novel ...

Tethered Bilayer Lipid Membranes to Monitor Heat Transfer between Gold Nanoparticles and Lipid Membranes

Here we report a protocol to investigate the heat transfer between irradiated gold nanoparticles (GNPs) and bilayer lipid membranes by electrochemistry using tethered bilayer lipid membranes (tBLMs) assembled on gold electrodes. Irradiated modified GNPs, such as streptavidin-conjugated GNPs, are embedded in tBLMs containing target molecules, such as biotin. By using this approach, the heat transfer processes between irradiated GNPs and model bilayer lipid membrane with entities of interest are mediated by a horizontally focused laser beam. The thermal predictive computational model is used to confirm the electrochemically induced conductance changes in the tBLMs. Under the specific conditions used, detecting heat pulses required specific attachment of the gold nanoparticles to the membrane surface, while unbound gold nanoparticles failed to elicit a measurable response. This technique serves as a powerful detection biosensor which can be directly utilized for the design and development of strategies for thermal therapies that permits optimization of the laser parameters, particle size, particle coatings and composition.

This work outlines a protocol to achieve dynamic, non-invasive monitoring of heat transfer from laser-irradiated gold nanoparticles to tBLMs. The system combines impedance spectroscopy for the real-time measurement of conductance changes across the tBLMs, with a horizontally focused laser beam that drives gold nanoparticle illumination, for heat production.

Here we report a protocol to investigate the heat transfer between irradiated gold nanoparticles (GNPs) and bilayer lipid membranes by electrochemistry ...

Optical Clearing of Plant Tissues for Fluorescence Imaging

It is challenging to directly observe the internal structure of multi-layered and opaque plant specimens, without dissection, under a microscope. In addition, autofluorescence attributed to chlorophyll hampers the observation of fluorescent proteins in plants. For a long time, various clearing reagents have been used to make plants transparent. However, conventional clearing reagents diminish fluorescent signals; therefore, it has not been possible to observe the cellular and intracellular structures with fluorescent proteins. Reagents were developed that can clear plant tissues by removing chlorophyll while maintaining fluorescent protein stability. A detailed protocol is provided here for the optical clearing of plant tissues using clearing reagents, ClearSee (CS) or ClearSeeAlpha (CSA). The preparation of cleared plant tissues involves three steps: fixation, washing, and clearing. Fixation is a crucial step in maintaining the cellular structures and intracellular stability of fluorescent proteins. The incubation time for clearing depends on the tissue type and species. In Arabidopsis thaliana, the time required for clearing with CS was 4 days for leaves and roots, 7 days for seedlings, and 1 month for pistils. CS also required a relatively short time of 4 days to make the gametophytic leaves of Physcomitrium patens transparent. In contrast, pistils in tobacco and torenia produced brown pigment due to oxidation during CS treatment. CSA reduced the brown pigment by preventing oxidation and could make tobacco and torenia pistils transparent, although it took a relatively long time (1 or 2 months). CS and CSA were also compatible with staining using chemical dyes, such as DAPI (4',6-diamidino-2-phenylindole) and Hoechst 33342 for DNA and Calcofluor White, SR2200, and Direct Red 23 for the cell wall. This method can be useful for whole-plant imaging to reveal intact morphology, developmental processes, plant-microbe interactions, and nematode infections.

Here, a method is described for making plant tissues transparent while maintaining the stability of fluorescent proteins. This technique facilitates deep imaging of cleared plant tissues without physical sectioning.

It is challenging to directly observe the internal structure of multi-layered and opaque plant specimens, without dissection, under a microscope. In ...