Research was supported by National Institute of Allergy and Infectious Diseases (NIH-NIAID) award R15AI079685.

1. Solution Preparation
NOTE: Prepare solutions described in steps 1.1&#8211;1.9 and 1.8&#8211;1.11 in order to produce E. coli spheroplasts and B. megaterium protoplasts, respectively.
<ol>
	<li>Prepare 1 M Tris-Cl, pH 7.8 by dissolving 10.34 g Tris HCl and 4.17 g of Tris OH in 50 mL of dH2O in a 125 mL flask. Sterilize by filtering through a 25 mm syringe filter with a 0.2 &#181;m membrane and store in a conical tube at room temperature.</li>
	<li>Prepare solution A (20 mM MgCl2, 0.7 M sucrose, 10 mM Tris-Cl, pH 7.8) by dissolving 0.10 g MgCl2 (95.2 g/mol) and 11.98 g sucrose (342.3 g/mol) in 25 mL dH2O in a 125 mL flask.Add 500 &#181;L 1 M Tris-Cl (pH 7.8) and adjust the volume to 50 mL. Sterilize by filtering through a 25 mm syringe filter with a 0.2 &#181;m membrane and store in a conical tube at room temperature.</li>
	<li>Prepare solution B (10 mM MgCl2, 0.8 M sucrose, 10 mM Tris-Cl, pH 7.8) by dissolving 0.05 g MgCl2 (95.2 g/mol) and 13.69 g sucrose (342.3 g/mol) in 25 mL dH2O in a 125 mL flask. Add 500 &#181;L 1 M Tris-Cl (pH 7.8) and adjust the volume to 50 mL. Sterilize by filtering through a 25 mm syringe filter with a 0.2 &#181;m membrane and store in a conical tube at room temperature.</li>
	<li>Prepare 0.8 M sucrose by dissolving 13.69 g sucrose (342.3 g/mol) in 50 mL dH2O in a 125 mL flask. Sterilize by filtering through a 25 mm syringe filter with a 0.2 &#181;m membrane and store in a conical tube at room temperature.</li>
	<li>Prepare 5 mg/mL deoxyribonuclease I (DNase I) by dissolving 0.015 g DNase I in 3 mL of dH2O in a 50 mL flask. Sterilize by filtering through a 25 mm syringe filter with a 0.2 &#181;m membrane. Aliquot solution into microfuge tubes and store at -20 &#176;C.</li>
	<li>Prepare 0.125 M ethylenediaminetetraacetic acid (EDTA), pH 8.0 by dissolving 0.698 g EDTA disodium dehydrate (372.2 g/mol) into 15 mL dH2O in a 125 mL flask. Sterilize by filtering through a 25 mm syringe filter with a 0.2 &#181;m membrane and store in a conical tube at room temperature.</li>
	<li>Prepare 600 &#181;g/mL cephalexin by dissolving 0.03 g of cephalexin hydrate (365.404 g/mol) in 50 mL of dH2O in a 125 mL flask. Sterilize by filtering through a 25 mm syringe filter with a 0.2 &#181;m membrane and store in a conical tube at 4 &#176;C.</li>
	<li>Prepare 5 mg/mL lysozyme by dissolving 0.015 g lysozyme in 3 mL of dH2O in a 50 mL flask. Sterilize by filtering through a 25 mm syringe filter with a 0.2 &#181;m membrane. Aliquot solution into microfuge tubes and store at -20 &#176;C.</li>
	<li>Prepare 3% w/v Trypticase Soy Broth (TSB) by dissolving 30 g of TSB in 1 L of dH2O in a 2 L flask. Aliquot the solution into flasks containing 25 mL or 100 mL TSB to be used in the preparation of E. coli spheroplasts or B. megaterium protoplasts, respectively. Autoclave flasks to sterilize the liquid medium. Sterile TSB can be stored at either room temperature or 4 &#176;C.</li>
	<li>Prepare solution C (1 M sucrose, 0.04 M maleate, 0.04 M MgCl2, pH 6.5) by dissolving 34.23 g sucrose (342.3 g/mol), 0.46 g maleic acid (116.07 g/mol), and 0.38 g MgCl2 (95.21 g/mol) in 100 mL of dH2O in a 250 mL flask. Adjust pH to 6.5. Sterilize by filtering through a 25 mm syringe filter with a 0.2 &#181;m membrane and store in a conical tube at room temperature.</li>
	<li>Prepare 200 mL protoplast medium by mixing 100 mL 3% w/v TSB and 100 mL solution C in a 1 L glass bottle. Autoclave to sterilize the solution and store at room temperature.</li>
</ol>
2. Preparation of Overnight Culture
NOTE: Perform sections 2-4 using appropriate sterile techniques. If desired, a bacterial strain can contain a plasmid for antibiotic resistance to reduce the potential contamination. If using a strain with antibiotic resistance, add the necessary antibiotics in steps 2.1, 3.1&#8211;3.2, and 4.1&#8211;4.4.
<ol>
	<li>Prepare an overnight culture by picking a single colony of bacteria using a sterile pipette tip and placing it into a 14 mL culture tube containing 2&#8211;3 mL of 3% w/v TSB. Incubate at 37 &#176;C, while shaking for 16&#8211;21 h.</li>
</ol>
3. Preparation of Gram-negative E. coli Spheroplasts
<ol>
	<li>In a 250 mL flask, dilute the overnight culture 1:100 in 25 mL volume of 3% w/v TSB and incubate the bacterial solution at 37 &#176;C, while shaking for approximately 2.5 h until the solution has reached an optical density of 0.5&#8211;0.8 at 600 nm. Measure the optical density using a spectrophotometer.</li>
	<li>In a 250 mL flask, dilute this culture 1:10 in 30 mL of 3% w/v TSB and incubate at 37 &#176;C, while shaking for 2.5 h in the presence of 60 &#181;g/mL cephalexin (347.4 g/mol) to produce single cell filaments of about 50&#8211;150 &#181;m in length, which are observable under 1,000X magnification using a light microscope (Figure 1B).</li>
	<li>Harvest the filaments by centrifuging the bacterial solution at 1,500 x g, 4 &#176;C for 4 min. Decant and discard the supernatant, reserving the pellet.</li>
	<li>Wash the filaments by gently adding 1 mL of 0.8 M sucrose, being careful not to disturb the pellet. Incubate for 1 min, and then discard the supernatant without disturbing the pellet.</li>
	<li>Add 150 &#181;L of 1 M Tris-Cl (pH 7.8), 120 &#181;L of 5 mg/mL lysozyme, 30 &#181;L of 5 mg/mL DNase I, and 120 &#181;L of 0.125 M EDTA in their respective order to the pellet and incubate the solution at room temperature for 10 min.</li>
	<li>Add 1 mL of solution A gradually over 1 min to the solution prepared in 3.5 using a micropipette while gently swirling the solution by hand. Incubate the solution for 4 min at room temperature.</li>
	<li>Put 7 mL of 4 &#176;C solution B into two 15 mL conical tubes. Add equal amounts of the solution prepared in 3.6 to each of these two tubes. Centrifuge the solution at 1,500 x g, 4 &#176;C for 4 min.</li>
	<li>Using a serological pipette, carefully remove all but 1-2 mL of supernatant without disturbing the pellet. Resuspend the pellet by gently pipetting up and down using a P1000 micropipette. Visually check to see spheroplasts formation by observing the sample at 1,000X magnification using a light microscope (Figure 1C).</li>
	<li>Store the spheroplasts at -20 &#176;C for up to a week or until they have gone through 3 freeze-thaw cycles.</li>
</ol>
4. Preparation of Gram-positive B. megaterium Protoplasts
<ol>
	<li>In a 250 mL flask, dilute the overnight culture 1:1,000 in 100 mL of 3% w/v TSB and incubate the bacterial solution at 37 &#176;C, while shaking for approximately 4.5 h until the solution has reached an optical density of 0.9-1.0 at 600 nm. Measure the optical density using a spectrophotometer.</li>
	<li>Pour the liquid culture into two 50 mL conical tubes and centrifuge at 2,000 x g, 4 &#176;C for 10 min.</li>
	<li>Using a serological pipette, discard the supernatant from both conical tubes. Resuspend each pellet in 2.5 mL protoplast medium and combine the resuspended solutions into a single conical tube. Pipette the combined resuspended solution in a 125 mL flask.</li>
	<li>Add 1 mL of 5 mg/mL lysozyme and incubate for 1 h at 37 &#176;C, while shaking.</li>
	<li>Monitor the growth of protoplasts under 1,000x magnification using a light microscope, noting any irregularities, such as bacteria that appear as swollen rods instead of spheres (Figure 2). Using a serological pipette, transfer the solution to a 15 mL conical tube and centrifuge at 2,000 x g, 4 &#176;C for 10 min.</li>
	<li>Decant the supernatant and re-suspend the pellet in 5 mL protoplast media. Store protoplasts at -20 &#176;C for up to a week or until they have gone through 3 freeze-thaw cycles.</li>
</ol>
5. Preparation of Peptide Solution and Membrane Dye for Imaging
<ol>
	<li>Peptide solution
	<ol>
		<li>In a microfuge tube wrapped in aluminum foil, dissolve 2 mg of FITC labeled BF2 P11A in 800 &#181;L dH2O. Use a peptide containing at least one tryptophan residue in order to conduct protein concentration measurements.</li>
		<li>Spectrophotometrically, measure the absorbance of the peptide solution at 280 nm in triplicate. Calculate the peptide concentration using the molar extinction coefficient for tryptophan (5,700/Mcm).</li>
		<li>Dilute the peptide concentration in dH2O to a final concentration of 100&#8211;200 &#181;M. Store in a microfuge tube at -20 &#176;C wrapped in aluminum foil to protect from light.</li>
	</ol>
	</li>
	<li>Membrane dye
	<ol>
		<li>In a microfuge tube, prepare a 10 mM stock of di-8-ANEPPS (592.9 g/mol) by dissolving 5 mg of di-8-ANEPPS in 843.3 &#181;L of DMSO. Store at 4 &#176;C wrapped in aluminum foil to protect from light.</li>
		<li>In a microfuge tube, prepare 1 mL of 0.03 mM di-8-ANEPPS by adding 3 &#181;L of 10 mM di-8-ANEPPS to 997 &#181;L DMSO. Store in a microfuge tube at 4 &#176;C wrapped in aluminum foil to protect from light.</li>
	</ol>
	</li>
</ol>
6. Visualization of E. coli Spheroplasts and B. megaterium Protoplasts Using Confocal Microscopy
<ol>
	<li>Pipette 5 &#181;L of spheroplasts or protoplasts onto a poly-L-lysine coated glass slide. Add 2 &#181;L of FITC labeled AMP (100-200 &#181;M) to the slide and incubate for 3 min protected from light.</li>
	<li>Add 1 &#181;L of the membrane dye di-8-ANEPPS (0.03 mM) to the slide and incubate for 3 min protected from light. Cover with a glass coverslip and seal with the nail polish.</li>
	<li>Turn on the confocal microscope and argon laser. Adjust the argon laser to 20% power output and 20% transmission.</li>
	<li>Set emission wavelength ranges of 499&#8211;532 nm and 670&#8211;745 nm for the FITC-labeled peptide emission channel and di-8-ANEPPS-labeled membrane dye emission channel, respectively.</li>
	<li>Image protoplasts or spheroplasts (Figure 1 and Figure 2) with a 63X objective. Use imaging software to obtain 8-bit, 512 x 512 composite z-stack images composed of 0.5 &#181;m slices of the entirety of the spheroplast or protoplast. 
	NOTE: Take careful consideration to reduce the emission bleed-through while imaging spheroplasts and protoplasts. See discussion for details.</li>
</ol>
7. Characterization of AMP Localization
<ol>
	<li>Open the composite z-stack image in imaging software. Locate the center-most slice of the spheroplast or protoplast and place one circular Region of Interest (ROI) (0.3 &#181;m in diameter) on the membrane (ROI 1), one at the center of the spheroplast or protoplast (ROI 2), and one away from spheroplast or protoplast to measure the background fluorescence (ROI 3) (Figure 5). Avoid the inclusion of saturated pixels. The fluorescence intensity in each ROI can be calculated by imaging software. 
	NOTE: In instances when peptide does not localize to the entirety of the membrane, draw ROI 1 on an area of the membrane where the peptide is localized.</li>
	<li>Use the following equation to determine the ratio of intracellular peptide fluorescence intensity to membrane peptide fluorescence intensity: 
	<img alt="Equation" src="/files/ftp_upload/57904/57904eq1.jpg" /> 
	NOTE: The fluorescence intensity in ROI 3 is subtracted from fluorescence intensities in ROI 2 and ROI 1 in order to account for background fluorescence.</li>
</ol>

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J., Adler, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fusion+of+bacterial+spheroplasts+by+electric+fields.">Fusion of bacterial spheroplasts by electric fields.</a> Biochim. Biophys. Acta. 819 (1), (1985).</li></ol>

No conflicts of interest are declared.

The protocols presented here make it feasible for researchers to more rapidly obtain larger sample sizes of bacterial images because the enlarged, spherical bacteria are much easier to locate, orient, and image. This enhanced ability to collect data is valuable in several respects. First, it enables a more systematic quantitative analysis of peptide localization patterns. While qualitative trends can be demonstrated from smaller sets of images, only a large sample set of high-quality images reveal more nuanced trends in ...

Antimicrobial peptides (AMPs) have gained attention due to their potential use as alternatives to conventional antibiotics1,2,3,4,5. AMPs kill bacteria by either translocating across the cell membrane and interacting with intracellular components such as nucleic acids or by permeabilizing the membrane causing leakage of cell contents6. In addition to their use as antibiotics, translocating AMPs may be adapted for drug delivery applications because they can non-disruptively cross the impermeable cell membrane7,8. We, therefore, seek to understand fundamental AMP mechanisms of action to lay the foundation for their use in drug design.
Confocal microscopy offers a way to assess localization patterns of fluorescently labeled AMPs in bacterial cells providing insights into their mechanism of action9,10,11,12,13,14. By labeling the membrane of the bacteria, one can determine if a fluorescently labeled peptide localizes to the membrane or the intracellular space of a bacterial cell. However, this technique is limited by the small size and rod shape of bacteria, which can make imaging challenging due to the resolution limits of conventional light microscopes and the variable orientation of the bacteria on the slide15.
The goal of the presented method is to enable enhanced visualization of the fluorescently labeled peptide localization patterns using confocal microscopy. Visualization is enhanced by turning the small, thin, rod-shaped gram-negative Escherichia coli (E. coli) and gram-positive Bacillus megaterium (B. megaterium) bacteria into enlarged, spherical forms referred to as spheroplasts (for gram-negative strains) and protoplasts (for gram-positive strains)16,17,18,19,20,21. Spheroplasts and protoplasts are easier to image because of both their increased size and their symmetric shape, which makes the orientation of a bacterium on a slide irrelevant for its imaging. In addition, we present a systematic approach to quantitatively analyze confocal microscopy data in order to characterize AMPs as either membrane localizing or translocating. Applying these methods makes it easier to distinguish fluorescently labeled peptide localization patterns. The protocols presented here can be used to assess the localization of a variety of membrane-active agents other than AMPs, including cell-penetrating peptides.
One distinct advantage of this technique is that it provides insights into the mechanism of action of AMPs on a single cell level, which may reveal cell-to-cell heterogeneity15, as opposed to other fluorescence assays commonly used to identify the mechanisms of action of AMPs, which only provide bulk estimates9,22,23,24,25. The use of spheroplasts and protoplasts in order to assess AMP cell entry is particular useful26&#160;because they are more physiologically relevant15&#160;than other models used for assessing cell entry, such as lipid vesicles24.

<table>
	<tbody>
		<tr>
			<td>Trizma hydrocloride (Tris HCl)</td>
			<td>Sigma</td>
			<td>T3253</td>
			<td></td>
		</tr>
		<tr>
			<td>Trizma base (Tris OH)</td>
			<td>Sigma</td>
			<td>T1503</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium chloride</td>
			<td>Sigma</td>
			<td>M8266</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma</td>
			<td>S7903</td>
			<td></td>
		</tr>
		<tr>
			<td>Lysozyme</td>
			<td>Sigma</td>
			<td>L6876</td>
			<td></td>
		</tr>
		<tr>
			<td>Deoxyribonuclease I</td>
			<td>Sigma</td>
			<td>D4527</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethylenediaminetetraacetic acid</td>
			<td>Sigma</td>
			<td>106361</td>
			<td>Used Sigma 106361 in original protocol development; 106361 discontinued with ED2SS as replacement</td>
		</tr>
		<tr>
			<td>Cephalexin hydrate</td>
			<td>Sigma</td>
			<td>C4895</td>
			<td></td>
		</tr>
		<tr>
			<td>Ampicillin</td>
			<td>Fisher Scientific</td>
			<td>BP1760</td>
			<td></td>
		</tr>
		<tr>
			<td>BBL Trypticase soy broth</td>
			<td>Fisher Scientific</td>
			<td>B11768</td>
			<td></td>
		</tr>
		<tr>
			<td>BF2 P11A FITC</td>
			<td>NeoScientific</td>
			<td>Custom ordered</td>
			<td></td>
		</tr>
		<tr>
			<td>di-8-ANEPPS</td>
			<td>Biotium</td>
			<td>61012</td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Sigma</td>
			<td>34869</td>
			<td>Used Sigma D8779 in original protocol development; D8779 discontinued with 34869 as replacement</td>
		</tr>
		<tr>
			<td>Maleic acid</td>
			<td>Sigma</td>
			<td>M0375</td>
			<td></td>
		</tr>
		<tr>
			<td>Acrodisc 25 mm Syringe Filter w/ 0.2 &#956;m HT Tuffryn Membrane</td>
			<td>Pall Corporation</td>
			<td>4192</td>
			<td></td>
		</tr>
		<tr>
			<td>Laser scanning confocal microscope</td>
			<td>Leica Microsystems</td>
			<td>TCS SP5 II</td>
			<td>For image acquisition</td>
		</tr>
		<tr>
			<td>Leica Application Suite, Advanced Fluorescence</td>
			<td>Leica Microsystems</td>
			<td></td>
			<td>For image processing</td>
		</tr>
	</tbody>
</table>

production and visualization of bacterial spheroplasts and protoplasts to characterize antimicrobial peptide localization

The use of confocal microscopy as a method to assess peptide localization patterns within bacteria is commonly inhibited by the resolution limits of conventional light microscopes. As the resolution for a given microscope cannot be easily enhanced, we present protocols to transform the small rod-shaped gram-negative Escherichia coli (E. coli) and gram-positive Bacillus megaterium (B. megaterium) into larger, easily imaged spherical forms called spheroplasts or protoplasts. This transformation allows observers to rapidly and clearly determine whether peptides lodge themselves into the bacterial membrane (i.e., membrane localizing) or cross the membrane to enter the cell (i.e., translocating). With this approach, we also present a systematic method to characterize peptides as membrane localizing or translocating. While this method can be used for a variety of membrane-active peptides and bacterial strains, we demonstrate the utility of this protocol by observing the interaction of Buforin II P11A (BF2 P11A), an antimicrobial peptide (AMP), with E. coli spheroplasts and B. megaterium protoplasts.

By enlarging bacteria and making them spherical, we can easily distinguish whether peptides localize to the bacterial membrane or readily translocate across the bacterial membrane. The resolution limits of conventional light microscopes make it challenging to distinguish whether peptide signals arise from the membrane or intracellular space in normal bacteria because signals localized to the membrane will appear to overlap with the intracellular space (Figure 3A</stro...

Watch this Scientific Journal Video about Production and Visualization of Bacterial Spheroplasts and Protoplasts to Characterize Antimicrobial Peptide Localization at JoVE.com

Production and Visualization of Bacterial Spheroplasts and Protoplasts to Characterize Antimicrobial Peptide Localization

Here we present a protocol to produce gram-negative Escherichia coli (E. coli) spheroplasts and gram-positive Bacillus megaterium (B. megaterium) protoplasts to clearly visualize and rapidly characterize peptide-bacteria interactions. This provides a systematic method to define membrane localizing and translocating peptides.

The use of confocal microscopy as a method to assess peptide localization patterns within bacteria is commonly inhibited by the resolution limits of ...

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JoVE Journal

Biology

11.6K Views.  Wellesley College. Here we present a protocol to produce gram-negative Escherichia coli (E. coli) spheroplasts and gram-positive Bacillus megaterium (B. megaterium) protoplasts to clearly visualize and rapidly characterize peptide-bacteria interactions. This provides a systematic method to define membrane localizing and translocating peptides.

Video: Production and Visualization of Bacterial Spheroplasts and Protoplasts to Characterize Antimicrobial Peptide Localization

Fluorescence Activated Cell Sorting of Plant Protoplasts

High-resolution, cell type-specific analysis of gene expression greatly enhances understanding of developmental regulation and responses to environmental stimuli in any multicellular organism. In situ hybridization and reporter gene visualization can to a limited extent be used to this end but for high resolution quantitative RT-PCR or high-throughput transcriptome-wide analysis the isolation of RNA from particular cell types is requisite. Cellular dissociation of tissue expressing a fluorescent protein marker in a specific cell type and subsequent Fluorescence Activated Cell Sorting (FACS) makes it possible to collect sufficient amounts of material for RNA extraction, cDNA synthesis/amplification and microarray analysis.
An extensive set of cell type-specific fluorescent reporter lines is available to the plant research community. In this case, two marker lines of the Arabidopsis thaliana root are used: PSCR::GFP (endodermis and quiescent center) and PWOX5::GFP (quiescent center). Large numbers (thousands) of seedlings are grown hydroponically or on agar plates and harvested to obtain enough root material for further analysis. Cellular dissociation of plant material is achieved by enzymatic digestion of the cell wall. This procedure makes use of high osmolarity-induced plasmolysis and commercially available cellulases, pectinases and hemicellulases to release protoplasts into solution.
FACS of GFP-positive cells makes use of the visualization of the green versus the red emission spectra of protoplasts excited by a 488 nm laser. GFP-positive protoplasts can be distinguished by their increased ratio of green to red emission. Protoplasts are typically sorted directly into RNA extraction buffer and stored for further processing at a later time.
This technique is revealed to be straightforward and practicable. Furthermore, it is shown that it can be used without difficulty to isolate sufficient numbers of cells for transcriptome analysis, even for very scarce cell types (e.g. quiescent center cells). Lastly, a growth setup for Arabidopsis seedlings is demonstrated that enables uncomplicated treatment of the plants prior to cell sorting (e.g. for the cell type-specific analysis of biotic or abiotic stress responses). Potential supplementary uses for FACS of plant protoplasts are discussed.

A method for isolating specific cell types from plant material is demonstrated. This technique employs transgenic marker lines expressing fluorescent proteins in particular cell types, cellular dissociation and Fluorescence Activated Cell Sorting. Additionally, a growth setup is established here that facilitates treatment of Arabidopsis thaliana seedlings prior to cell sorting.

High-resolution, cell type-specific analysis of gene expression greatly enhances understanding of developmental regulation and responses to environmental ...

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 ...

Nano-fEM: Protein Localization Using Photo-activated Localization Microscopy and Electron Microscopy

Mapping the distribution of proteins is essential for understanding the function of proteins in a cell. Fluorescence microscopy is extensively used for protein localization, but subcellular context is often absent in fluorescence images. Immuno-electron microscopy, on the other hand, can localize proteins, but the technique is limited by a lack of compatible antibodies, poor preservation of morphology and because most antigens are not exposed to the specimen surface. Correlative approaches can acquire the fluorescence image from a whole cell first, either from immuno-fluorescence or genetically tagged proteins. The sample is then fixed and embedded for electron microscopy, and the images are correlated 1-3. However, the low-resolution fluorescence image and the lack of fiducial markers preclude the precise localization of proteins. 
Alternatively, fluorescence imaging can be done after preserving the specimen in plastic. In this approach, the block is sectioned, and fluorescence images and electron micrographs of the same section are correlated 4-7. However, the diffraction limit of light in the correlated image obscures the locations of individual molecules, and the fluorescence often extends beyond the boundary of the cell. 
Nano-resolution fluorescence electron microscopy (nano-fEM) is designed to localize proteins at nano-scale by imaging the same sections using photo-activated localization microscopy (PALM) and electron microscopy. PALM overcomes the diffraction limit by imaging individual fluorescent proteins and subsequently mapping the centroid of each fluorescent spot 8-10. 
We outline the nano-fEM technique in five steps. First, the sample is fixed and embedded using conditions that preserve the fluorescence of tagged proteins. Second, the resin blocks are sectioned into ultrathin segments (70-80 nm) that are mounted on a cover glass. Third, fluorescence is imaged in these sections using the Zeiss PALM microscope. Fourth, electron dense structures are imaged in these same sections using a scanning electron microscope. Fifth, the fluorescence and electron micrographs are aligned using gold particles as fiducial markers. In summary, the subcellular localization of fluorescently tagged proteins can be determined at nanometer resolution in approximately one week.

We describe a method to localize fluorescently tagged proteins in electron micrographs. Fluorescence is first localized using photo-activated localization microscopy on ultrathin sections. These images are then aligned to electron micrographs of the same section.

Mapping the distribution of proteins is essential for understanding the function of proteins in a cell. Fluorescence microscopy is extensively used for ...

Measurement and Analysis of Extracellular Acid Production to Determine Glycolytic Rate

Extracellular measurement of oxygen consumption and acid production is a simple and powerful way to monitor rates of respiration and glycolysis1. Both mitochondrial (respiration) and non-mitochondrial (other redox) reactions consume oxygen, but these reactions can be easily distinguished by chemical inhibition of mitochondrial respiration. However, while mitochondrial oxygen consumption is an unambiguous and direct measurement of respiration rate2, the same is not true for extracellular acid production and its relationship to glycolytic rate 3-6. Extracellular acid produced by cells is derived from both lactate, produced by anaerobic glycolysis, and CO2, produced in the citric acid cycle during respiration. For glycolysis, the conversion of glucose to lactate- + H+ and the export of products into the assay medium is the source of glycolytic acidification. For respiration, the export of CO2, hydration to H2CO3 and dissociation to HCO3- + H+ is the source of respiratory acidification. The proportions of glycolytic and respiratory acidification depend on the experimental conditions, including cell type and substrate(s) provided, and can range from nearly 100% glycolytic acidification to nearly 100% respiratory acidification 6. Here, we demonstrate the data collection and calculation methods needed to determine respiratory and glycolytic contributions to total extracellular acidification by whole cells in culture using C2C12 myoblast cells as a model.

Glycolysis is a defining metabolic marker in multiple biological systems. Monitoring glycolysis by measuring the extracellular flux of H+ is common, but requires correction to be quantitative and unambiguous. Here, we demonstrate how to gather and correct extracellular flux data to distinguish between respiratory and glycolytic sources of extracellular acidification.

Extracellular measurement of oxygen consumption and acid production is a simple and powerful way to monitor rates of respiration and glycolysis1. Both ...

Targeted RNA Sequencing Assay to Characterize Gene Expression and Genomic Alterations

RNA sequencing (RNAseq) is a versatile method that can be utilized to detect and characterize gene expression, mutations, gene fusions, and noncoding RNAs. Standard RNAseq requires 30 - 100 million sequencing reads and can include multiple RNA products such as mRNA and noncoding RNAs. We demonstrate how targeted RNAseq (capture) permits a focused study on selected RNA products using a desktop sequencer. RNAseq capture can characterize unannotated, low, or transiently expressed transcripts that may otherwise be missed using traditional RNAseq methods. Here we describe the extraction of RNA from cell lines, ribosomal RNA depletion, cDNA synthesis, preparation of barcoded libraries, hybridization and capture of targeted transcripts and multiplex sequencing on a desktop sequencer. We also outline the computational analysis pipeline, which includes quality control assessment, alignment, fusion detection, gene expression quantification and identification of single nucleotide variants. This assay allows for targeted transcript sequencing to characterize gene expression, gene fusions, and mutations.

We describe a targeted RNA sequencing-based method that includes preparation of indexed cDNA libraries, hybridization and capture with custom probes and data analysis to interrogate selected transcripts for gene expression, mutations, and gene fusions. Targeted RNAseq permits cost-effective, rapid evaluation of selected transcripts on a desktop sequencer.

RNA sequencing (RNAseq) is a versatile method that can be utilized to detect and characterize gene expression, mutations, gene fusions, and noncoding ...

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 ...

Utilizing pHluorin-tagged Receptors to Monitor Subcellular Localization and Trafficking

Understanding membrane protein trafficking, assembly, and expression requires an approach that differentiates between those residing in intracellular organelles and those localized on the plasma membrane. Traditional fluorescence-based measurements lack the capability to distinguish membrane proteins residing in different organelles. Cutting edge methodologies transcend traditional methods by coupling pH-sensitive fluorophores with total internal reflection fluorescence microscopy (TIRFM). TIRF illumination excites the sample up to approximately 150 nm from the glass-sample interface, thus decreasing background, increasing the signal to noise ratio, and enhancing resolution. The excitation volume in TIRFM encompasses the plasma membrane and nearby organelles such as the peripheral ER. Superecliptic pHluorin (SEP) is a pH sensitive version of GFP. Genetically encoding SEP into the extracellular domain of a membrane protein of interest positions the fluorophore on the luminal side of the ER and in the extracellular region of the cell. SEP is fluorescent when the pH is greater than 6, but remains in an off state at lower pH values. Therefore, receptors tagged with SEP fluoresce when residing in the endoplasmic reticulum (ER) or upon insertion in the plasma membrane (PM) but not when confined to a trafficking vesicle or other organelles such as the Golgi. The extracellular pH can be adjusted to dictate the fluorescence of receptors on the plasma membrane. The difference in fluorescence between TIRF images at neutral and acidic extracellular pH for the same cell corresponds to a relative number of receptors on the plasma membrane. This allows a simultaneous measurement of intracellular and plasma membrane resident receptors. Single vesicle insertion events can also be measured when the extracellular pH is neutral, corresponding to a low pH trafficking vesicle fusing with the plasma membrane and transitioning into a fluorescent state. This versatile technique can be exploited to study localization, expression, and trafficking of membrane proteins.

Labeling the extracellular domain of a membrane protein with a pH sensitive fluorophore, superecliptic pHluorin (SEP), allows subcellular localization, expression, and trafficking to be determined. Imaging SEP-labeled proteins with total internal reflection fluorescence microscopy (TIRFM) enables the quantification of protein levels in the peripheral ER and plasma membrane.

Understanding membrane protein trafficking, assembly, and expression requires an approach that differentiates between those residing in intracellular ...

Measuring Endoreduplication by Flow Cytometry of Isolated Tuber Protoplasts

Endoreduplication, the replication of a cell's nuclear genome without subsequent cytokinesis, yields cells with increased DNA content and is associated with specialization, development and increase in cellular size. In plants, endoreduplication seems to facilitate the growth and expansion of certain tissues and organs. Among them is the tuber of potato (Solanum tuberosum), which undergoes considerable cellular expansion in fulfilling its function of carbohydrate storage. Thus, endoreduplication may play an important role in how tubers are able to accommodate this abundance of carbon. However, the cellular debris resulting from crude nuclear isolation methods of tubers, methods that can be used effectively with leaves, precludes the estimation of the tuber endoreduplication index (EI). This article presents a technique for assessing tuber endoreduplication through the isolation of protoplasts while demonstrating representative results obtained from different genotypes and compartmentalized tuber tissues. The major limitations of the protocol are the time and reagent costs required for sample preparation as well as relatively short lifespan of samples after lysis of protoplasts. While the protocol is sensitive to technical variation, it represents an improvement over traditional methods of nuclear isolation from these large specialized cells. Possibilities for improvements to the protocol such as recycling enzyme, the use of fixatives, and other alterations are proposed.

The protocol described herein is a method for measuring endoreduplication within tubers of potato (Solanum tuberosum). It includes plasmolysis and protoplast extraction steps to decrease the noise and debris in downstream flow cytometric analysis.

Endoreduplication, the replication of a cell's nuclear genome without subsequent cytokinesis, yields cells with increased DNA content and is associated ...

A Simple Method for Isolation of Soybean Protoplasts and Application to Transient Gene Expression Analyses

Soybean (Glycine max (L.) Merr.) is an important crop species and has become a legume model for the studies of genetic and biochemical pathways. Therefore, it is important to establish an efficient transient gene expression system in soybean. Here, we report a simple protocol for the preparation of soybean protoplasts and its application for transient functional analyses. We found that young unifoliate leaves from soybean seedlings resulted in large quantities of high quality protoplasts. By optimizing a PEG-calcium-mediated transformation method, we achieved high transformation efficiency using soybean unifoliate protoplasts. This system provides an efficient and versatile model for examination of complex regulatory and signaling mechanisms in live soybean cells and may help to better understand diverse cellular, developmental and physiological processes of legumes.

We developed a simple and efficient protocol for the preparation of large quantities of soybean protoplasts to study complex regulatory and signaling mechanisms in live cells.

Soybean (Glycine max (L.) Merr.) is an important crop species and has become a legume model for the studies of genetic and biochemical pathways. ...

Assessment of Zebrafish Lens Nucleus Localization and Sutural Integrity

The zebrafish is uniquely suited to genetic manipulation and in vivo imaging, making it an increasingly popular model for reverse genetic studies and for generation of transgenics for in vivo imaging. These unique capabilities make the zebrafish an ideal platform to study ocular lens development and physiology. Our recent findings that an Aquaporin-0, Aqp0a, is required for stability of the anterior lens suture, as well as for the shift of the lens nucleus to the lens center with age led us to develop tools especially suited to analyzing the properties of zebrafish lenses. Here we outline detailed methods for lens dissection that can be applied to both larval and adult lenses, to prepare them for histological analysis, immunohistochemistry and imaging. We focus on analysis of lens suture integrity and cortical cell morphology and compare data generated from dissected lenses with data obtained from in vivo imaging of lens morphology made possible by a novel transgenic zebrafish line with a genetically encoded fluorescent marker. Analysis of dissected lenses perpendicular to their optical axis allows quantification of the relative position of the lens nucleus along the anterior-posterior axis. Movement of the lens nucleus from an initial anterior position to the center is required for normal lens optics in adult zebrafish. Thus, a quantitative measure of lens nuclear position directly correlates with its optical properties.

These protocols were developed to analyze cortical lens morphology, structural integrity of the zebrafish lens sutures in fixed and live lenses and to measure the position of the zebrafish lens nucleus along the anterior-posterior axis.

The zebrafish is uniquely suited to genetic manipulation and in vivo imaging, making it an increasingly popular model for reverse genetic studies and for ...