We would like to thank Gero Miesenb&#246;ck and Tim Ryan for sharing pHluorin cDNA used to generate reagents in this study, Nathan Wright, Joanna Poprawski and Lydia Nyasae for excellent technical assistance, members of the Wendland lab for helpful discussions, and Michael McCaffery and Erin Pryce at the Integrated Imaging Center (Johns Hopkins) for advice and assistance with microscopy and flow cytometry. This work was supported by grants from the National Institutes of Health (to B.W., GM60979) and the National Science Foundation (to B.W., MCB 1024818). K.W. was supported in part by a training grant from the National Institutes of Health (T32-GM007231). A.F.O. was supported by developmental funds from the Department of Biological Sciences at Duquesne University and National Science Foundation CAREER grant 553143.

1. Solutions and Media
<ol>
	<li>Prepare the Following Stocks, Media and Plates:
	<ol>
		<li>To prepare 10x YNB, dissolve 67 g of yeast nitrogen base lacking amino acids in 1 L of water. Sterilize by filtration through a 0.22 &#181;m nitrocellulose filter.</li>
		<li>Prepare YNB medium using 1x YNB (from 10x stock) with 2% dextrose (from a sterile 50% w/v stock) and 1x amino acid/nutrient mixture (from 100x stock, see step 1.1.4). For plasmid selection, omit individual amino acids or nutrients as needed. For induction of Mup1 expression, additionally omit methionine from the amino acid/nutrient mixture.</li>
		<li>Prepare YNB plates using 1x YNB (from 10x stock), 2% dextrose (from a sterile 50% w/v stock), 0.7 g/L amino acid/nutrient mixture (see step 1.1.5) and 2.5% agar. Prepare plate medium by autoclaving 25 g of agar and 0.7 g of amino acid/nutrient mixture per 860 ml of water. Allow the mixture to cool to 55 &#176;C, then add 100 ml of 10x YNB and 40 ml of 50% glucose. Pour plates (approximately 30 ml of medium per 10 cm dish), allow the medium to solidify at room temperature, and store plates at 4 &#176;C.</li>
		<li>Prepare 100x amino acid/nutrient stock solution (for liquid medium) by dissolving the following in 100 ml of deionized water: 0.1 g L-Methionine, 0.3 g each of L-leucine and L-lysine, and 0.2 g each of L-histidine, L-tryptophan, adenine and uracil (other amino acids and nutrients may be included as needed for specific strains). Use gentle heat if needed to dissolve, and sterilize through a 0.45 &#181;m nitrocellulose filter. Store at room temperature with protection from light. For plasmid selection, prepare stock solutions lacking specific components as needed.</li>
		<li>Prepare amino acid/nutrient mixture (for plates) by combining the following: 0.5 g of adenine, 4 g of L-leucine, and 2 g each of L-histidine, L-lysine, L-methionine, L-tryptophan, L-tyrosine and uracil. Grind with a mortar and pestle, and store with protection from light. For plasmid selection, prepare mixtures lacking specific components as needed, and use 0.7 g of amino acid mixture per liter of plate medium (see step 1.1.3).</li>
	</ol>
	</li>
	<li>Prepare 8-well chamber slides for microscopy by coating the surface of the slide with concanavalin A (ConA). To each well of the chamber slide, add 25 &#181;l of 2 mg/ml ConA dissolved in water. Using a pipette tip, spread the ConA across the bottom surface of the well, then allow the slide to air dry at room temperature for at least 4-8 hr. Ideally, use chamber slides within 24 hr of ConA coating.</li>
	<li>Generate yeast strains with in-frame fusions of GFP or pHluorin to the endocytic cargo of interest using the polymerase chain reaction (PCR)-based integration method described in Longtine et al. and Goldstein and McCusker25,26. 
	NOTE: Plasmids containing pHluorin tagging cassettes with resistance genes for kanamycin (KANMX6) and nourseothricin (NATMX4) were described previously20.
	<ol>
		<li>Amplify GFP or pHluorin cassettes containing selectable markers with primers containing additional sequences specific to the site of genomic integration20,25.</li>
		<li>Transform the resulting PCR product into the desired yeast strain using the lithium acetate method27.</li>
		<li>Select for integration of the cassette by growing cells on YPD plates containing the appropriate drug. To prepare plates, autoclave 10 g yeast extract, 20 g peptone, 0.1 g tryptophan and 25 g agar in 960 ml water. Before pouring plates, allow the mixture to cool to 55 &#176;C, then add 40 ml of 50% glucose and either G418 to a final concentration of 200 &#181;g/ml for KANMX6 selection, or nourseothricin to a final concentration of 100 &#181;g/ml for NATMX4 selection.</li>
		<li>Confirm integration of the GFP or pHluorin tag in individual colonies by PCR and/or Western blotting using anti-GFP antibodies, as well as by detection of the tagged protein by fluorescence microscopy20. 
		NOTE: Specific yeast strains and plasmids used in this study are listed in Tables 1 and 2, respectively.</li>
	</ol>
	</li>
</ol>
2. Endpoint Assay for Steady-state Fluorescence of Constitutively Internalized Endocytic Cargo
<ol>
	<li>Inoculate cells (~3 small colonies) in 5 ml of YNB medium lacking amino acids or nutrients as required for plasmid maintenance (if applicable). Grow cells for 16-24 hr in an orbital shaking incubator at 30 &#176;C, with 250 rpm shaking. Alternatively, streak a ~1-2 mm colony of cells onto a YNB plate lacking amino acids or nutrients for plasmid selection, and grow cells overnight at 30 &#176;C.</li>
	<li>Measure the density (OD600) of each overnight culture using a spectrophotometer. Prepare a 5 ml dilution at 0.35-0.4 OD600/ml in YNB medium lacking amino acids or nutrients for plasmid maintenance, and grow for approximately 3 hr at 30 &#176;C with shaking. If using cells streaked on a plate, skip this step and proceed with step 2.4 (see below).</li>
	<li>Measure the density (OD600) of cells, which should now be between 0.6-1.0 OD600/ml. Transfer 1.5 ml of culture to a microfuge tube, and pellet cells at 8,000 rpm (6,800 x g) for 2 min. Remove 1,475 &#181;l of supernatant.</li>
	<li>Bring cells to a fluorescence microscope. Before imaging each sample, prepare a slide by resuspending the cells in the remaining medium, and mount 3 &#181;l of cell suspension under a cover slip. Alternatively, prepare 8-well chamber slides as described below in protocol 3, using YNB medium as appropriate for plasmid selection. Ideally, image cells with a 100X, 1.4 or higher numerical aperture (NA) oil immersion objective lens, a 12- or 16-bit camera, and excitation/emission filters optimized for GFP fluorescence. 
	NOTE: If using cells grown by streaking on a plate, prepare the slide by placing 3 &#181;l of fresh YNB medium on a cover slip. Using a pipette tip, collect a small colony of cells from the plate, disperse the cells into the YNB medium, and mount the resulting suspension on a glass slide immediately before imaging.</li>
	<li>Image 4-6 random fields of cells for each condition, using the same acquisition parameters (i.e., filter sets and exposure time) for all images within an experiment. 
	NOTE: Collecting a brightfield or DIC image for each field can be helpful for cells where the pHluorin signal is quenched in MVB and vacuolar compartments.</li>
	<li>Quantify fluorescence intensity of at least 30-50 cells for each condition (see protocol 5).</li>
</ol>
3. Kinetic Assay for Quantification of Endocytosis of Cargos that Undergo Regulated Endocytosis
<ol>
	<li>Inoculate 2-3 colonies of yeast cells (approximately 2 mm in diameter per colony) in 5 ml of YNB medium lacking methionine and additional amino acids or nutrients for maintaining plasmid selection. Grow cultures for 16-24 hr at 30 &#176;C with shaking. 
	NOTE: In this paper, protocols for studying regulated endocytic cargo proteins will make use of the methionine permease, Mup1. Other cargos that undergo regulated endocytosis are also suitable for tagging with pHluorin (for example, the uracil permease Fur4, which accumulates at the plasma membrane in the absence of uracil, and internalizes in response to externally applied uracil); for these cargos, media conditions should be modified to reflect the specific expression, induction and internalization conditions for that cargo.</li>
	<li>Measure the density (OD600) of each overnight culture approximately 3 hr before imaging. Prepare a 5 ml dilution at 0.35-0.4 OD600/ml in YNB medium lacking methionine and additional amino acids or nutrients for plasmid maintenance, and grow at 30 &#176;C with shaking.</li>
	<li>Pre-equilibrate the microscope&#39;s environmental chamber or heated stage (if available) to 30 &#176;C during the 3 hr incubation of step 3.2.</li>
	<li>Transfer 1 ml of cell culture to a microfuge tube, and pellet cells at 8,000 rpm (6,800 x g) for 2 min. Remove 975 &#181;l of supernatant.</li>
	<li>Prepare a ConA-treated, 8-well glass-bottomed chamber slide for imaging (protocol 1.2).</li>
	<li>Add 200 &#181;l of YNB medium lacking methionine and additional amino acids or nutrients for plasmid selection to each well. Resuspend the cell pellet from step 3.4 in the remaining supernatant, and drop 2.5 &#181;l of cell suspension into the center of each well. Disperse cells across the surface of the well by pipetting up and down, and allow cells to settle for 5-10 min.</li>
	<li>Place the chamber slide on an inverted fluorescence microscope equilibrated to 30 &#176;C (see step 3.3). Locate a field of cells using brightfield illumination.</li>
	<li>Add 50 &#181;l of YNB medium containing 100 &#181;g/ml methionine (pre-warmed to 30 &#176;C) to the 200 &#181;l of medium in the well to obtain a final methionine concentration of 20 &#181;g/ml. Avoid disrupting the medium in the well in order to prevent the cells from floating from the bottom.</li>
	<li>Allow the cells to equilibrate for 2 min before beginning imaging. Immediately before capturing the first image, adjust the microscope to the desired focal plane (typically, an equatorial focal plane works best).</li>
	<li>Acquire GFP and brightfield (or DIC) images at 5 min intervals for 45-60 min, using identical acquisition parameters for all images within a time series (for example, 100X 1.4 NA oil immersion objective and 500 msec acquisition time using GFP filter, 100 msec acquisition time using DIC filter). 
	NOTE: If the microscope&#39;s stage and imaging software do not allow for automatic correction of focal plane drift, it may be necessary to manually readjust the focus prior to each imaging time point. Moreover, acquisition times will vary between microscopes due to differences in illumination and filters, and optimal conditions should be determined manually prior to beginning the experiment. If an endocytic cargo other than Mup1 is used, it may be necessary to modify the acquisition time and imaging intervals, as well as the duration of the experiment, to reflect the internalization kinetics of that cargo.</li>
	<li>Quantify fluorescence intensity of individual cells at each time point (see protocol 5), and express values as a percentage of the initial intensity of the first image.</li>
</ol>
4. Endpoint Assay for Quantification of Endocytic Cargos that Undergo Regulated Endocytosis
<ol>
	<li>Prepare cells for imaging as described in protocol 3 (steps 3.1 through 3.6).</li>
	<li>Image 4-5 random fields of cells for each condition immediately before adding methionine, using brightfield and GFP filter sets.</li>
	<li>Add 50 &#181;l of YNB medium containing 100 &#181;g/ml methionine (pre-warmed to 30 &#176;C) to the 200 &#181;l of medium in the well to obtain a final methionine concentration of 20 &#181;g/ml. Avoid disrupting the medium in the well in order to prevent the cells from floating from the bottom.</li>
	<li>Image 4-5 random fields of cells 30 min after addition of methionine, using the same acquisition parameters as for the 0 min time point (step 4.2).</li>
	<li>Quantify fluorescence intensity of at least 30-50 cells for each time point (see protocol 5). Express values as the percentage of Mup1-pHluorin internalized after 30 min using the formula: [Mean intensity at 0 min - Mean intensity at 30 min]/[Mean intensity at 0 min] x 100%. 
	NOTE: It is possible to simultaneously perform up to eight endpoints assays by staggering the start time of each assay. Table 3 provides an example workflow for eight simultaneous assays.</li>
</ol>
5. Post-acquisition Analysis
<ol>
	<li>Export images from acquisition software as a 16-bit tagged-image file format (.tif or .tiff format). 
	NOTE: Other file formats may also be suitable, and should ideally use lossless compression algorithms. 8-bit images are generally not suitable for quantification purposes.</li>
	<li>Open files using ImageJ software (freely available at &#39;Open&#39; option in the &#39;File&#39; menu. Use the fluorescence image for quantification, and the brightfield/DIC image as a reference to identify the location of cells and to exclude dead cells (if present), which appear darker than live cells when viewed by DIC and are autofluorescent when viewed with GFP excitation/emission filters.</li>
	<li>Perform a background subtraction on the fluorescence image. For images with an uneven background signal, use the &#39;Background Subtraction&#39; algorithm found in the &#39;Process menu&#39;. Use the default setting (rolling ball radius of 50 pixels), which is generally sufficient for quantification purposes. 
	NOTE: An alternative background subtraction method for images with uniform background is described in step 5.3.1.-5.3.2; however, the above method also works for uniform background.
	<ol>
		<li>For images with uniform background (i.e., background intensity is approximately constant at all edges and in the center of the image), perform a manual background subtraction. Click on the &#39;freehand selections&#39; button found in the main ImageJ window, and select 3-5 random regions within the image that do not contain cells.</li>
		<li>Open the &#39;Set Measurements&#39; function located in the &#39;Analyze&#39; menu, select the &#39;Mean Gray Value&#39; parameter, and measure intensity values using the &#39;Measure&#39; function (also found in the &#39;Analyze&#39; menu). Calculate the average value from the 3-5 measured regions, and subtract from all subsequent measurements in that image.</li>
	</ol>
	</li>
	<li>For quantification of kinetic assays (protocol 3), generate a single, stacked image for the time series. Open the fluorescence images from each time point in chronological order, and generate a stack using the &#39;Images to Stack&#39; function (found in the &#39;Image&#39; menu, under the Stacks sub-menu).</li>
	<li>Outline a cell using the freehand selection tool, making sure to include the entire cell but as little area outside of the cell as possible.</li>
	<li>Measure the following parameters: &#39;Area, Integrated Density&#39; and &#39;Mean Gray Value&#39;. Select parameters using the &#39;Set Measurements&#39; function, located in the &#39;Analyze&#39; menu. 
	NOTE: Integrated Density corresponds to the sum of all pixel intensities within the selected region, and Mean Gray Value corresponds to [Integrated Density/Area], which corrects intensity values for cell size.</li>
	<li>Open the &#39;ROI Manager&#39; located in the &#39;Analyze&#39; menu, under the &#39;Tools&#39; sub-menu. In the ROI Manager window, click the &#39;Add&#39; button to enter the highlighted region as a new region of interest (ROI).</li>
	<li>Repeat steps 5.4 through 5.6 for the remaining cells in the image. Exclude any dead cells as assessed by brightfield/DIC and fluorescence (dead cells appear darker in DIC images, and have autofluorescent cytoplasmic signal when viewed with GFP excitation/emission; see Figure 2F), and any cells that touch the edge of the image. Save the selected ROIs as a .zip file by clicking the &#39;More&#39; button in the &#39;ROI Manager&#39; window, and export all measurements to a spreadsheet or statistical analysis software.</li>
	<li>For quantification of kinetic assays (protocol 3), outline and measure a cell from the initial time point as described in steps 5.4 and 5.5. Use the same ROI for measurement of all subsequent time points in the experiment.</li>
	<li>For endpoint assays (protocols 2 and 4), measure a minimum of 30-50 cells for each condition. Perform quantification for at least 2-3 images, and include all cells in each image that meet the criteria outlined in step 5.8 in the analysis.</li>
	<li>Assess statistical significance between samples using one-way ANOVA, followed by Tukey&#39;s multiple comparison test for post hoc analysis.</li>
</ol>
6. Population Analysis of Mup1-pHluorin Endocytosis by Flow Cytometry
<ol>
	<li>Inoculate 2-3 colonies of yeast cells (approximately 2 mm in diameter per colony) in 0.5 ml of YNB medium lacking methionine and additional amino acids or nutrients for maintaining plasmid selection. Grow cells in 5 ml round-bottom tubes for 16-24 hr at 30 &#176;C on a roller drum.</li>
	<li>Add 1.5 ml of YNB medium lacking methionine and additional amino acids or nutrients for plasmid maintenance, and grow at 30 &#176;C for an additional 3 hr on a roller drum.</li>
	<li>Transfer 1 ml of cells into each of two 5 ml round-bottom tubes. Add 0.25 ml of YNB -Met medium to the first tube (-Met condition), and 0.25 ml of YNB medium containing 100 &#181;g/ml methionine to the second tube to give a final methionine concentration of 20 &#181;g/ml (+Met condition). Incubate cells at 30 &#176;C for 45 min on a roller drum.</li>
	<li>Analyze cells by flow cytometry, selecting forward scatter (FS) as a measure of cell size, and Mup1-pHluorin fluorescence intensity from a fluorescein isothiocyanate (FITC) filter as a measure of cargo internalization.</li>
	<li>Use the slider buttons to adjust the voltage of the FS and FITC channels to optimize detection for the size range and fluorescence intensity of the yeast cells being used (for these experiments, settings used were 50 V for FS and 400 V for FITC). 
	NOTE: Voltage for both channels will vary depending on the flow cytometer being used. This step should be done before or during the 45 min treatment with methionine (step 6.3).</li>
	<li>Measure FS and fluorescence intensity for 10,000 cells from each condition, using the high flow rate setting. Generate a scatter plot of FS against fluorescence intensity: click &#39;add graph&#39;, select FITC (x-axis) and FS (y-axis), and graph 1,000 points (the software will display values for 1,000 cells, even though 10,000 cells were measured). 
	NOTE: For maximal consistency, analyze cells 45-50 min after addition of methionine; for experiments involving large numbers of samples, stagger addition of methionine to accommodate the time required for flow cytometry analysis.</li>
	<li>Include -Met and +Met conditions for wild-type (WT) cells as a control. Using the gate function, assign a vertical gate using the WT +Met condition, such that approximately 5% of the brightest cells fall to the right of the gate. Use this gating for all other conditions in the experiment.</li>
</ol>

The authors declare no competing interests.

Applications of pHluorin for quantification of endocytic events in yeast cells makes use of transmembrane cargos in which the fluorescent tag is fused to the cytoplasmic tail of the protein (Figure 1A). For the assays described here, cargos are brightly fluorescent when the pHluorin tag is exposed to neutral environments, but become quenched when the pHluorin tag encounters acidic conditions. Thus, a pHluorin-tagged endocytic cargo is readily detected at the cytoplasmic face of the plasma membrane and on...

Vesicular trafficking plays an important role in maintaining organelle identity and function in eukaryotic cells, and is a key mechanism for regulating protein and membrane composition of individual cellular compartments. At the plasma membrane, fusion of exocytic vesicles delivers new proteins and membranes to the surface of the cell, whereas vesicles generated through endocytosis remove membrane and proteins from the surface for subsequent recycling or targeting to the lysosome. Thus, endocytosis is important for nutrient uptake and for responses to the extracellular environment. Exocytosis and endocytosis are balanced to regulate plasma membrane surface area, and to allow turnover of damaged proteins.
Studies in yeast and mammalian cells have identified a large number of proteins involved in endocytosis, as well as multiple endocytic pathways that promote internalization of specific cargos or that act in response to a variety of environmental conditions. The best-studied pathway is clathrin-mediated endocytosis (CME), in which clathrin and cytosolic accessory proteins assemble into a coat structure to stabilize the nascent endocytic vesicle. Experiments in the budding yeast Saccharomyces cerevisiae have yielded key insights into the dynamics and order of recruitment for many endocytic proteins1-3. Notably, the CME machinery is highly conserved through evolution such that the majority of CME-related proteins in yeast have human orthologs; thus, budding yeast has been an important tool for understanding mechanisms of endocytosis that are conserved in higher eukaryotes. For example, studies using budding yeast determined that formation and maturation of CME structures (referred to as cortical actin patches in yeast) involves the sequential recruitment of many proteins, beginning with clathrin and cargo-binding adaptor proteins, followed by recruitment of additional endocytic accessory proteins, activation of Arp2/3-mediated actin polymerization, and recruitment of proteins involved in vesicle scission1,2. Recruitment of CME machinery proteins to cortical actin patches is a highly ordered and stereotypical process, and the precise order of recruitment for many proteins has been established with respect to other components of the CME machinery. Importantly, recent studies have confirmed a similar order of recruitment for CME proteins at clathrin-coated pits in mammalian cells4.
In addition to CME, many cell types possess one or more clathrin-independent endocytic (CIE) pathways that rely on alternative mechanisms to promote vesicle formation and internalization from the plasma membrane5-7. In yeast, we recently identified a CIE pathway that utilizes the small GTPase Rho1 and its activating guanine nucleotide exchange factor (GEF), Rom18,9. Rho1 activates the formin Bni1 to promote actin polymerization10,11, which is required for this form of clathrin-independent endocytosis in yeast12. Moreover, the &#945;-arrestin family of proteins recruit the ubiquitin ligase Rsp5 to promote cargo ubiquitination and subsequent internalization through CME13-17, and also promote cargo internalization via the CIE pathway, possibly through direct interaction with proteins involved in CIE18. A variety of CIE pathways also exist in mammalian cells, including clathrin-independent and phagocytic pathways that rely on the Rho1 ortholog, RhoA9,19. The role of RhoA in mammalian CIE is poorly understood; thus, studies in budding yeast may provide additional mechanistic insights that are applicable to mammalian CIE.
Accurate quantification of endocytic events can provide important information about the roles of specific proteins in regulating endocytosis, and can reveal the effects of mutations on cargo internalization or progression through the endocytic pathway. For this purpose, biochemical methods can be utilized to monitor ligand uptake or to measure rates of degradation of endocytic cargo proteins. In living cells, fusion of green fluorescent protein (GFP) and its variants to cargos of interest allows direct visualization of cargo transport. However, GFP-tagged cargos are of limited use for quantification of endocytosis because GFP is resistant to degradation in the lysosome (or vacuole in yeast). Moreover, GFP fluorescence is only partially sensitive to changes in pH, and remains detectable within the vacuole lumen20,21. Consequently, fluorescence of the GFP tag persists in the vacuole long after the remainder of the cargo has been degraded, and whole cell-based quantification of cargo intensity may be inaccurate due to long-term GFP fluorescence in the vacuole.
In order to overcome the drawbacks of vacuolar GFP fluorescence, we previously made use of superecliptic pHluorin, a pH-sensitive variant of GFP that fluoresces brightly at neutral pH, but loses fluorescence in acidic environments such as the lumen of the vacuole/lysosome20,22,23. Placing a pHluorin tag on the cytoplasmic tail of endocytic cargos permits visualization of the cargos at the plasma membrane and on early endosomes, where the pHluorin tag remains exposed to the cytoplasm (Figure 1A). As early endosomes mature, the endosomal sorting complex required for transport (ESCRT) machinery packages surface-localized cargos into vesicles that bud into the lumen of the endosome, generating multivesicular bodies (MVBs)24. For cargos that have been incorporated into internal MVB vesicles, the pHluorin tag faces toward the vesicle lumen. Internal MVB vesicles are acidified; thus, pHluorin-tagged cargos lose fluorescence on early endosomes as they mature into MVBs20. Subsequent fusion of the MVB with the vacuole delivers the MVB luminal contents for degradation, and pHluorin tags remain quenched in the acidic environment of the vacuole lumen.
This paper provides detailed descriptions of quantitative endocytic assays using cargos with cytoplasmic pHluorin tags in yeast. Strains expressing pHluorin-tagged cargos can be used in kinetic and/or endpoint assays, depending on the properties and trafficking behavior of the specific cargo. In addition, some pHluorin-tagged cargos are amenable to high-throughput analysis methods, including flow cytometry. Importantly, the pHluorin tag is a versatile tool for studying endocytic events in living cells, allowing quantification of endocytosis and comparison of endocytic function in wild-type and mutant strains.

<table><tbody><tr><td>Adenine</td><td>Sigma</td><td>A8626-25G</td><td>Use for preparation of amino acid mixture and stock solution</td></tr><tr><td>Bacto-agar</td><td>Fisher</td><td>BP1423-2</td><td>Use for preparation of plate media</td></tr><tr><td>BD Difco Yeast Nitrogen Base (without amino acids)</td><td>BD</td><td>291920</td><td>Use for preparation of liquid and plate media</td></tr><tr><td>Concanavalin A</td><td>Sigma</td><td>C5275-5MG</td><td>Use for coating of chamber slides</td></tr><tr><td>Dextrose</td><td>Fisher</td><td>BP350-1</td><td>Use for preparation of liquid and plate media</td></tr><tr><td>L-Histidine</td><td>Fisher</td><td>BP382-100</td><td>Use for preparation of amino acid mixture and stock solution</td></tr><tr><td>L-Leucine</td><td>Acros</td><td>125121000</td><td>Use for preparation of amino acid mixture and stock solution</td></tr><tr><td>L-Lysine</td><td>Fisher</td><td>BP386-100</td><td>Use for preparation of amino acid mixture and stock solution</td></tr><tr><td>L-Methionine</td><td>Fisher</td><td>BP388-100</td><td>Use for preparation of amino acid mixture and stock solution</td></tr><tr><td>L-Tryptophan</td><td>Fisher</td><td>BP395-100</td><td>Use for preparation of amino acid mixture and stock solution</td></tr><tr><td>L-Tyrosine</td><td>Acros</td><td>140641000</td><td>Use for preparation of amino acid mixture</td></tr><tr><td>Nunc Lab-Tek Chambered Coverglass (8-well)</td><td>Thermo Scientific</td><td>155411</td><td>Use for kinetic and endpoint assays of Mup1-pHluorin internalization</td></tr><tr><td>Uracil</td><td>Sigma</td><td>U0750-100G</td><td>Use for preparation of amino acid mixture and stock solution</td></tr><tr><td>Axiovert 200 inverted microscope</td><td>Carl Zeiss</td><td>Custom Build</td><td></td></tr><tr><td>100X/1.4 Plan-Apochromat Oil Immersion Objective Lens</td><td>Carl Zeiss</td><td></td><td>Objective should be 100X, 1.4NA or higher</td></tr><tr><td>Sensicam</td><td>Cooke Corporation</td><td></td><td>Camera should have 12-bit or higher dynamic range</td></tr><tr><td>X-Cite 120PC Q Illumination Source</td><td>Excelitas Technologies</td><td></td><td></td></tr><tr><td>Slidebook 5 software</td><td>Intelligent Imaging Innovations</td><td></td><td></td></tr><tr><td>ImageJ software</td><td>National Institutes of Health</td><td></td><td>http://imagej.nih.gov/ij/</td></tr></tbody></table>

applications of phluorin for quantitative, kinetic and high-throughput analysis of endocytosis in budding yeast

Green fluorescent protein (GFP) and its variants are widely used tools for studying protein localization and dynamics of events such as cytoskeletal remodeling and vesicular trafficking in living cells. Quantitative methodologies using chimeric GFP fusions have been developed for many applications; however, GFP is somewhat resistant to proteolysis, thus its fluorescence persists in the lysosome/vacuole, which can impede quantification of cargo trafficking in the endocytic pathway. An alternative method for quantifying endocytosis and post-endocytic trafficking events makes use of superecliptic pHluorin, a pH-sensitive variant of GFP that is quenched in acidic environments. Chimeric fusion of pHluorin to the cytoplasmic tail of transmembrane cargo proteins results in a dampening of fluorescence upon incorporation of the cargo into multivesicular bodies (MVBs) and delivery to the lysosome/vacuole lumen. Thus, quenching of vacuolar fluorescence facilitates quantification of endocytosis and early events in the endocytic pathway. This paper describes methods using pHluorin-tagged cargos for quantification of endocytosis via fluorescence microscopy, as well as population-based assays using flow cytometry.

Steady-state localization and quantification of the constitutively internalized endocytic cargo protein Ste3
To demonstrate that pHluorin-tagged cargos can be utilized for quantification of endocytosis in living cells, localization of chimeric GFP and pHluorin fusions with the cytoplasmic C-terminal tail of Ste3, the a-factor pheromone receptor in yeast, were compared. Ste3 is a G protein-coupled receptor (GPCR)...

Watch this Scientific Journal Video about Applications of pHluorin for Quantitative, Kinetic and High-throughput Analysis of Endocytosis in Budding Yeast at JoVE.com

Applications of pHluorin for Quantitative, Kinetic and High-throughput Analysis of Endocytosis in Budding Yeast

Accurate quantification of vesicular trafficking events often provides key insights into roles for specific proteins and the effects of mutations. This paper presents methods for using superecliptic pHluorin, a pH-sensitive GFP variant, as a tool for quantification of endocytic events in living cells using quantitative fluorescence microscopy and flow cytometry.

Green fluorescent protein (GFP) and its variants are widely used tools for studying protein localization and dynamics of events such as cytoskeletal ...

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JoVE Journal

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10.8K Views.  The Johns Hopkins University. Accurate quantification of vesicular trafficking events often provides key insights into roles for specific proteins and the effects of mutations. This paper presents methods for using superecliptic pHluorin, a pH-sensitive GFP variant, as a tool for quantification of endocytic events in living cells using quantitative fluorescence microscopy and flow cytometry.

Video: Applications of pHluorin for Quantitative, Kinetic and High-throughput Analysis of Endocytosis in Budding Yeast

Single Molecule Fluorescence In Situ Hybridization (smFISH) Analysis in Budding Yeast Vegetative Growth and Meiosis

Single molecule fluorescence in situ hybridization (smFISH) is a powerful technique to study gene expression in single cells due to its ability to detect and count individual RNA molecules. Complementary to deep sequencing-based methods, smFISH provides information about the cell-to-cell variation in transcript abundance and the subcellular localization of a given RNA. Recently, we have used smFISH to study the expression of the gene&#160;NDC80&#160;during meiosis in budding yeast, in which two transcript isoforms exist and the short transcript isoform has its entire sequence shared with the long isoform. To confidently identify each transcript isoform, we optimized known smFISH protocols and obtained high consistency and quality of smFISH data for the samples acquired during budding yeast meiosis. Here, we describe this optimized protocol, the criteria that we use to determine whether high quality of smFISH data is obtained, and some tips for implementing this protocol in&#160;other yeast strains and growth conditions.

Single Molecule Fluorescence In Situ Hybridization smFISH Analysis in Budding Yeast Vegetative Growth and Meiosis

This single molecule fluorescence in situ hybridization protocol is optimized to quantify the number of RNA molecules in budding yeast during vegetative growth and meiosis.

Single molecule fluorescence in situ hybridization (smFISH) is a powerful technique to study gene expression in single cells due to its ability to detect ...

Genetics

Automated Quantification of Synaptic Fluorescence in C. elegans

Synapse strength refers to the amplitude of postsynaptic responses to presynaptic neurotransmitter release events, and has a major impact on overall neural circuit function. Synapse strength critically depends on the abundance of neurotransmitter receptors clustered at synaptic sites on the postsynaptic membrane. Receptor levels are established developmentally, and can be altered by receptor trafficking between surface-localized, subsynaptic, and intracellular pools, representing important mechanisms of synaptic plasticity and neuromodulation. Rigorous methods to quantify synaptically-localized neurotransmitter receptor abundance are essential to study synaptic development and plasticity. Fluorescence microscopy is an optimal approach because it preserves spatial information, distinguishing synaptic from non-synaptic pools, and discriminating among receptor populations localized to different types of synapses. The genetic model organism Caenorhabditis elegans is particularly well suited for these studies due to the small size and relative simplicity of its nervous system, its transparency, and the availability of powerful genetic techniques, allowing examination of native synapses in intact animals.
Here we present a method for quantifying fluorescently-labeled synaptic neurotransmitter receptors in C. elegans. Its key feature is the automated identification and analysis of individual synapses in three dimensions in multi-plane confocal microscope output files, tabulating position, volume, fluorescence intensity, and total fluorescence for each synapse. This approach has two principal advantages over manual analysis of z-plane projections of confocal data. First, because every plane of the confocal data set is included, no data are lost through z-plane projection, typically based on pixel intensity averages or maxima. Second, identification of synapses is automated, but can be inspected by the experimenter as the data analysis proceeds, allowing fast and accurate extraction of data from large numbers of synapses. Hundreds to thousands of synapses per sample can easily be obtained, producing large data sets to maximize statistical power. Considerations for preparing C. elegans for analysis, and performing confocal imaging to minimize variability between animals within treatment groups are also discussed. Although developed to analyze C. elegans postsynaptic receptors, this method is generally useful for any type of synaptically-localized protein, or indeed, any fluorescence signal that is localized to discrete clusters, puncta, or organelles.
The procedure is performed in three steps: 1) preparation of samples, 2) confocal imaging, and 3) image analysis. Steps 1 and 2 are specific to C. elegans, while step 3 is generally applicable to any punctate fluorescence signal in confocal micrographs.

Automated Quantification of Synaptic Fluorescence in C. elegans

The abundance of neurotransmitter receptors clustered at synapses strongly influences synaptic strength. This method quantifies fluorescently-labeled neurotransmitter receptors in three dimensions with single-synapse resolution in C. elegans, allowing hundreds of synapses to be rapidly characterized within a single sample without distortions introduced by z-plane projection.

Synapse strength refers to the amplitude of postsynaptic responses to presynaptic neurotransmitter release events, and has a major impact on overall ...

Neuroscience

High Throughput Fluorometric Technique for Assessment of Macrophage Phagocytosis and Actin Polymerization

The goal of fluorometric analysis is to serve as an efficient, cost effective, high throughput method of analyzing phagocytosis and other cellular processes. This technique can be used on a variety of cell types, both adherent and non-adherent, to examine a variety of cellular properties. When studying phagocytosis, fluorometric technique utilizes phagocytic cell types such as macrophages, and fluorescently labeled opsonized particles whose fluorescence can be extinguished in the presence of trypan blue. Following plating of adherent macrophages in 96-well plates, fluorescent particles (green or red) are administered and cells are allowed to phagocytose for varied amounts of time. Following internalization of fluorescent particles, cells are washed with trypan blue, which facilitates extinction of fluorescent signal from bacteria which are not internalized, or are merely adhering to the cell surface. Following the trypan wash, cells are washed with PBS, fixed, and stained with DAPI (nuclear blue fluorescent label), which serves to label nuclei of cells. By a simple fluorometric quantification through plate reading of nuclear (blue) or particle (red/green) fluorescence we can examine the ratio of relative fluorescence units of green:blue and determine a phagocytic index indicative of amount of fluorescent bacteria internalized per cell. The duration of assay using a 96-well method and multichannel pipettes for washing, from end of phagocytosis to end of data acquisition, is less than 45 min. Flow cytometry could be used in a similar manner but the advantage of fluorometry is its high throughput, rapid method of assessment with minimal manipulation of samples and quick quantification of fluorescent intensity per cell. Similar strategies can be applied to non adherent cells, live labeled bacteria, actin polymerization, and essentially any process utilizing fluorescence. Therefore, fluorometry is a promising method for its low cost, high throughput capabilities in the study of cellular processes.

Here we present a protocol to quantify phagocytosis of fluorescent particles by adherent macrophage cell line using a fluorometric method. This method facilitates a high throughput quantification of particle internalization as well as the resulting actin polymerization.

The goal of fluorometric analysis is to serve as an efficient, cost effective, high throughput method of analyzing phagocytosis and other cellular ...

Immunology and Infection

Use of pHluorin to Assess the Dynamics of Axon Guidance Receptors in Cell Culture and in the Chick Embryo

During development, axon guidance receptors play a crucial role in regulating axons sensitivity to both attractive and repulsive cues. Indeed, activation of the guidance receptors is the first step of the signaling mechanisms allowing axon tips, the growth cones, to respond to the ligands. As such, the modulation of their availability at the cell surface is one of the mechanisms that participate in setting the growth cone sensitivity. We describe here a method to precisely visualize the spatio-temporal cell surface dynamics of an axon guidance receptor both in vitro and in vivo in the developing chick spinal cord. We took advantage of the pH-dependent fluorescence property of a green fluorescent protein (GFP) variant to specifically detect the fraction of the axon guidance receptor that is addressed to the plasma membrane. We first describe the in vitro validation of such pH-dependent constructs and we further detail their use in vivo, in the chick spinal chord, to assess the spatio-temporal dynamics of the axon guidance receptor of interest.

We describe here the use of a pH-sensitive green fluorescent protein variant, pHluorin, to study the spatio-temporal dynamics of axon guidance receptors trafficking at the cell surface.&#160;The pHluorin-tagged receptor is expressed both in cell culture and in vivo, using electroporation of the chick embryo.

During development, axon guidance receptors play a crucial role in regulating axons sensitivity to both attractive and repulsive cues. Indeed, activation ...

Analysis of the Development of a Morphological Phenotype as a Function of Protein Concentration in Budding Yeast

Gene deletion and protein overexpression are common methods for studying functions of proteins. In this article, we describe a protocol for analysis of phenotype development as a function of protein concentration at population and single-cell levels in Saccharomyces cerevisiae. Although this protocol is based on the overexpression of a protein, it can easily be adapted for morphological phenotypes dependent on suppression of protein expression. Our lab is interested in studying the signaling properties of the endocytic adaptor protein epsin. To that purpose we used a dominant negative approach in which we over-expressed the conserved Epsin N-Terminal Homology (ENTH) domain in order to interfere with the functions of endogenous epsin-2 (Ent2 or YLR206W). We observed that overexpression of the ENTH domain of Ent2 (ENTH2) in wild type cells led to a cell division defect that is dependent on the mislocalization of a family of scaffolding proteins, septins.

Gene deletion and protein overexpression are common methods for studying functions of proteins. In this article, we describe a protocol for analysis of phenotype development as a function of protein concentration at population and single-cell levels in Saccharomyces cerevisiae.

Gene deletion and protein overexpression are common methods for studying functions of proteins. In this article, we describe a protocol for analysis of ...

Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using fluorochromes like Green Fluorescent Protein (GFP) directly attached to the protein of interest has become increasingly popular 1. Using GFP and similar fluorochromes the subcellular localisations and movements of proteins can be detected in a fluorescent microscope. Moreover, also the subnuclear localisation of a certain region of a chromosome can be studied using this technique. GFP is fused to the Lac Repressor protein (LacR) and ectopically expressed in the cell where tandem repeats of the lacO sequence has been inserted into the region of interest on the chromosome2. The LacR-GFP will bind to the lacO repeats and that area of the genome will be visible as a green dot in the fluorescence microscope. Yeast is especially suited for this type of manipulation since homologous recombination is very efficient and thereby enables targeted integration of the lacO repeats and engineered fusion proteins with GFP 3. Here we describe a quantitative method for live cell analysis of fission yeast. Additional protocols for live cell analysis of fission yeast can be found, for example on how to make a movie of the meiotic chromosomal behaviour 4. In this particular experiment we focus on subnuclear organisation and how it is affected during gene induction. We have labelled a gene cluster, named Chr1, by the introduction of lacO binding sites in the vicinity of the genes. The gene cluster is enriched for genes that are induced early during nitrogen starvation of fission yeast 5. In the strain the nuclear membrane (NM) is labelled by the attachment of mCherry to the NM protein Cut11 giving rise to a red fluorescent signal. The Spindle Pole body (SPB) compound Sid4 is fused to Red Fluorescent Protein (Sid4-mRFP) 6. In vegetatively growing yeast cells the centromeres are always attached to the SPB that is embedded in the NM 7. The SPB is identified as a large round structure in the NM. By imaging before and 20 minutes after depletion of the nitrogen source we can determine the distance between the gene cluster (GFP) and the NM/SPB. The mean or median distances before and after nitrogen depletion are compared and we can thus quantify whether or not there is a shift in subcellular localisation of the gene cluster after nitrogen depletion.

The fission yeast, Schizosaccharomyces pombe, is a good model system to study basic cellular processes. Here we describe a method to perform quantitative live cell analysis of fission yeast. In this particular experiment we focus on organisation of the genome within the cell nucleus, but the method can also be used to study cytosolic factors.

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using ...

Acquiring Fluorescence Time-lapse Movies of Budding Yeast and Analyzing Single-cell Dynamics using GRAFTS

Fluorescence time-lapse microscopy has become a powerful tool in the study of many biological processes at the single-cell level. In particular, movies depicting the temporal dependence of gene expression provide insight into the dynamics of its regulation; however, there are many technical challenges to obtaining and analyzing fluorescence movies of single cells. We describe here a simple protocol using a commercially available microfluidic culture device to generate such data, and a MATLAB-based, graphical user interface (GUI) -based software package to quantify the fluorescence images. The software segments and tracks cells, enables the user to visually curate errors in the data, and automatically assigns lineage and division times. The GUI further analyzes the time series to produce whole cell traces as well as their first and second time derivatives. While the software was designed for S. cerevisiae, its modularity and versatility should allow it to serve as a platform for studying other cell types with few modifications.

We present a simple protocol to obtain fluorescence microscopy movies of growing yeast cells, and a GUI-based software package to extract single-cell time series data. The analysis includes automated lineage and division time assignment integrated with visual inspection and manual curation of tracked data.

Fluorescence time-lapse microscopy has become a powerful tool in the study of many biological processes at the single-cell level. In particular, movies ...

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 ...

High-resolution Imaging and Analysis of Individual Astral Microtubule Dynamics in Budding Yeast

Dynamic microtubules are fundamental to many cellular processes, and accurate measurements of microtubule dynamics can provide insight into how cells regulate these processes and how genetic mutations impact regulation. The quantification of microtubule dynamics in metazoan models has a number of associated challenges, including a high microtubule density and limitations on genetic manipulations. In contrast, the budding yeast model offers advantages that overcome these challenges. This protocol describes a method to measure the dynamics of single microtubules in living yeast cells. Cells expressing fluorescently tagged tubulin are adhered to assembled slide chambers, allowing for stable time-lapse image acquisition. A detailed guide for high-speed, four-dimensional image acquisition is also provided, as well as a protocol for quantifying the properties of dynamic microtubules in confocal image stacks. This method, combined with conventional yeast genetics, provides an approach that is uniquely suited for quantitatively assessing the effects of microtubule regulators or mutations that alter the activity of tubulin subunits.

Budding yeast is an advantageous model for studying microtubule dynamics in vivo due to its powerful genetics and the simplicity of its microtubule cytoskeleton. The following protocol describes how to transform and culture yeast cells, acquire confocal microscopy images, and quantitatively analyze microtubule dynamics in living yeast cells.

Dynamic microtubules are fundamental to many cellular processes, and accurate measurements of microtubule dynamics can provide insight into how cells ...

Application of Membrane and Cell Wall Selective Fluorescent Dyes for Live-Cell Imaging of Filamentous Fungi

The application of membrane and cell wall selective fluorescent dyes for live-cell imaging analyses of organelle dynamics in fungal cells started two decades ago and since then continues to contribute greatly to our understanding of the filamentous fungal lifestyle. This paper provides a practical guide for the utilization of the two membrane dyes FM 1-43 and FM 4-64 and the four cell wall stains Calcofluor White M2R, Solophenyl Flavine 7GFE 500, Pontamine Fast Scarlet 48 and Congo Red. The focus is on their low-dose application to ascertain artefact-free staining, their co-imaging properties, and their quantitative evaluation. The presented methods are applicable to all filamentous fungal samples that can be prepared in the described ways. The fundamental staining approaches can serve as starting points for adaptations to species that might require different cultivation conditions. First, biophysical and biochemical properties are reviewed as their understanding is essential for using these dyes as truly vital fluorescent stains. Secondly, step-by-step protocols are presented that detail the preparation of various fungal sample types for fluorescent live-cell imaging. Finally, example experiments illustrate different approaches to: (1) identify defects in the spatio-temporal organization of endocytosis in genetic mutants, (2) comparatively characterize shared and distinct co-localization of GFP-labeled target proteins in the endocytic pathway, (3) identify morphogenetic cell wall defects in a genetic mutant, and (4) monitor cell wall biogenesis in real time.

Vital fluorescent dyes are essential tools for live-cell imaging analyses in modern fungal cell biology. This paper details the application of established and lesser-known fluorescent dyes for tracking plasma membrane dynamics, endo-/exocytosis and cell wall morphogenesis in filamentous fungi.

The application of membrane and cell wall selective fluorescent dyes for live-cell imaging analyses of organelle dynamics in fungal cells started two ...

Applications of pHluorin for Quantitative, Kinetic and High-throughput Analysis of Endocytosis in Budding Yeast

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