This work was supported by NIH R01 GM50322 to P.M. Kane. The authors thank Dr. Rajini Rao, Johns Hopkins University for providing the yeast pHluorin plasmids and for advice on ratiometric pH measurements, and Dr. Gloria A. Martinez Munoz for working out these protocols for our lab.

1. Measurement of Vacuolar pH In Vivo Using BCECF-AM
<ol>
	<li>Grow a 50 ml liquid culture of the yeast strain to be measured in the desired medium overnight. The goal is to have cells in mid-log phase (OD600 (optical density at 600 nm) measurement of approximately 0.8 for the suspension).</li>
	<li>Pellet the yeast cells by centrifugation. Resuspend the pellet in 0.6 ml of the growth medium and transfer to a microcentrifuge tube that has been weighed previously. Pellet the cells again in a microcentrifuge at 2,000 x g for 60 sec. Remove the supernatant as completely as possible and then weigh the cell pellet. Resuspend the pellet to a final density of 0.5 g/ml (w/v); a culture of this volume will give a cell pellet of approximately 200 mg, and 200 &mu;l of buffer would be added to give a final volume of 400 &mu;l.</li>
	<li>Add BCECF-AM to the cell suspension at a final concentration of 50 mM from a 12 mM stock prepared in DMSO. Mix well and then incubate the cells at 30 &deg;C for 30 min. on a rocking platform or roller drum.</li>
	<li>While the cells are incubating, prepare calibration buffers. The calibration buffer contains 50 mM MES (2-(N-morpholino)ethanesulfonic acid), 50 mM HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid), 50 mM KCl, 50 mM NaCl, 0.2 M ammonium acetate, 10 mM sodium azide, and 10 mM 2-deoxyglucose, and is adjusted to the pH values appropriate for your calibration range with NaOH or HCl. (Caution: Sodium azide is highly toxic and should be handled with care.) For measurement of pH in wild-type cells, we would prepare several calibration mixtures at pH 5.5 to 6.5, but for mutants expected to have a more alkaline vacuolar pH (vma mutants), we would prepare additional, higher pH buffers.</li>
	<li>Aliquot 2 ml of calibration buffer for each pH into 15 ml conical tubes, and add monensin (15 mM stock) and nigericin (2 mM stock) to give final concentrations of 110 &mu;M monensin and 15 &mu;M nigericin, respectively. Mix well on a vortex mixer. (Caution: Both monensin and nigericin are toxic and should be handled with care.)</li>
	<li>When the BCECF-AM incubation time is completed, pellet the cell suspension by centrifugation for 30 sec at 2,000 x g in a microcentrifuge. Resuspend the cells is 1 ml of growth medium lacking glucose and centrifuge as above. Repeat this wash step once, and then resuspend the pellet in 200 &mu;l of growth medium without glucose. Place on ice.</li>
	<li>Add 20 &mu;l of the cell suspension to each pH calibration tube prepared above. Incubate at 30 &deg;C for 30-60 min. on a roller drum.</li>
	<li>Set the fluorimeter to alternately measure at excitation wavelengths 450 nm and 490 nM, both with an emission wavelength of 535 nm. Set the sample chamber temperature to 30 &deg;C. Perform all measurements with continuous stirring of the mixture in the cuvette.</li>
	<li>Add 1.96 ml of 1 mM MES (adjusted to pH 5 or 7 depending on the pH of the cells' growth medium and the experimental design). Add 20 &mu;l of cell suspension into the cuvette. Initiate fluorescence measurements. We collect both continuous kinetic and time-point data in different experiments. For continuous kinetic data, we take measurements every 6 sec for 5 min, then add glucose to a final glucose concentration of 50 mM, and continue measurement for 5-10 min. For single time-point measurements, we generally take measurements at 1 and 5 min. after addition of cells to the cuvette, add glucose as in the kinetic measurements, and then take another measurement 5 min. after glucose addition. These additions can be varied, and additional components (inhibitors, etc) can also be added.</li>
	<li>After the experimental measurements are complete, remove the calibration tubes from the 30 &deg;C incubation and transfer the entire 2 ml volume for each to the fluorimeter cuvette. Measure fluorescence at the same settings (see 1.8) every 5 sec over a total of 30 sec for each sample.</li>
	<li>Export fluorescence data to Microsoft Excel. (For our fluorimeter, this requires export of data as text in tab-delimited form and import into Excel.) Obtain a calibration curve by calculating the ratio of fluorescence at 490 nm to 450 nm for each calibration mixture. The fluorescence ratio is then plotted vs. pH to get a calibration curve.</li>
	<li>Convert the experimental data to fluorescence ratio and calculate pH using the standard curve. Vacuolar pH can then plotted against time (kinetics of pH changes) or compared under various conditions (Figure 1) or in different mutant strains.</li>
</ol>
2. Measurement of Cytosolic pH In Vivo Using Yeast pHluorin
<ol>
	<li>Transform the desired yeast strain with the yeast pHluorin plasmid by standard protocols, and select for transformants on supplemented minimal medium lacking uracil (SC-uracil).</li>
	<li>Grow a 50 ml liquid culture of transformed cells in SC-uracil to mid-log phase (OD600 = 0.8 or lower).</li>
	<li>Prepare calibration standards as described for vacuolar pH, but buffer to a pH range appropriate for cytosolic pH measurements, generally pH 6-8. Aliquot 2 ml of calibration buffer adjusted to the desired pH into several 15 ml conical tubes. Add monensin and nigericin as described above (1.4) and mix well.</li>
	<li>Harvest the cells by centrifugation as described above, and resuspend the pellet in 600 &mu;l of the growth medium. Transfer to a weighed microcentrifuge tube, and pellet cells by centrifugation at 5,000 rpm for 30 sec. Resuspend the pellet in 1 ml of growth medium without glucose (SC-uracil, -glucose), pellet cells again and repeat. After the final centrifugation, remove the supernatant as thoroughly as possible and weigh the cell pellet. Resuspend cells to a final density of 0.5 g/ml in SC-uracil with no glucose.</li>
	<li>Add 20 &mu;l of cell suspension to each tube of calibration buffer and mix well on a vortex mixer. Incubate at 30 &deg;C on a spinning drum rotor for 60 min.</li>
	<li>Set up the fluorimeter for excitation at wavelengths 405 and 485 nm and an emission wavelength of 508 nm. Proceed with single time point or kinetic measurements and addition of glucose as described above for BCECF measurement.</li>
	<li>Measure fluorescence of calibration samples, and construct a calibration curve as for BCECF (see 1.10-1.11). A plot of fluorescence ratio vs. pH is linear over the range of most cytosolic pH measurements (pH 6.0-8.0) 9, so experimental fluorescence ratio measurements are easily converted to pH.</li>
</ol>

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The authors have no conflicts of interest to disclose.

We have utilized these protocols to address a number of aspects of pH homeostasis. For example, we have compared cytosolic and pH responses of wild-type and V-ATPase-deficient mutant cells 4,5. We have also examined the effects of altered growth conditions, particularly extracellular pH, on vacuolar pH response to glucose 3. Importantly, the responses we observe are both consistent with other methods of quantitative pH measurement and with biochemical data describing altered activities of the ...

pH homeostasis is a dynamic and highly regulated process in all organisms 1,2. Biochemical processes are tightly regulated by pH, and intracellular environments are tuned to narrow pH ranges to allow optimal activity of the resident enzymes. However, intracellular pH homeostasis can be challenged by rapid changes in environmental pH, metabolic shifts, and certain signaling pathways. In addition, intracellular pH can itself serve as an important signal. Finally, many organelles maintain lumenal pH values that are distinct from the surrounding cytosol and essential for organelle-specific functions.
The yeast Saccharomyces cerevisiae shares a number of pH homeostasis mechanisms with higher eukaryotes 2. In the acidic organelles of the endocytic/lysosomal pathway, pH is primarily controlled by the highly conserved vacuolar proton-translocating ATPase (V-ATPase), acting in tandem with many exchangers dependent on the pH gradient. All eukaryotic cells also have proton export mechanisms. In fungi and plants, a second, distinct proton pump at the plasma membrane, Pma1, exports metabolic protons and is believed to be the major determinant of cytosolic pH and plasma membrane potential. The genetic flexibility of S. cerevisiae and its commercial importance, have made it a very interesting and important model for studying pH homeostasis 2.
In addition to being the primary drivers of organelle acidification, V-ATPases are highly regulated enzymes and our lab is interested in understanding mechanisms of V-ATPase regulation. Towards this goal, we have been using in vivo pH measurements of vacuolar and cytosolic pH: 1) to monitor responses to changing extracellular conditions, such as glucose deprivation and readdition, 2) to examine the effects of mutations that compromise V-ATPase activity, and 3) to explore the coordination of organelle and plasma membrane proton pumps 3-5. These experiments only became possible through the development of robust ratiometric pH indicators amenable to use in yeast cells. Plant et al. first showed that BCECF (2'7'-Bis-(2-carboxyethyl)-5-(and 6)-carboxyfluorescein), which has been used widely to measure cytosolic pH in mammalian cells, accumulates in the yeast vacuole instead of the cytosol 6. This difference in BCECF localization has been attributed to the many hydrolytic enzymes in the vacuole, which are likely responsible for cleavage of the acetoxy methyl ester from BCECF-AM (acetoxymethyl ester of BCECF) and vacuolar retention 6. Ali et al. 7 further developed vacuolar pH measurement using BCECF and adapted these measurements to a fluorescence plate reader format. Brett et al. introduced yeast pHluorin as a means of measuring cytosolic pH in yeast by expressing a plasmid-borne ratiometric pH-sensitive GFP 8 under control of a yeast-specific promoter 9.
The excitation spectra of both BCECF and yeast pHluorin are sensitive to pH, so they are used as ratiometric pH indicators in which the ratio of fluorescence at two excitation wavelengths, measured at a single emission wavelength, provides a measure of pH 8,10. These yeast vacuolar and cytosolic pH sensors have been used for both single-cell and population-based measurements. Single-cell measurements 6,11 are performed by fluorescence microscopy and image analysis. Vacuolar or cytosolic fluorescence at the two wavelengths is measured for each cell. The population-based measurements are performed in either a microplate reader with appropriate fluorescence capabilities or in a fluorimeter. We have generally done our measurements in a fluorimeter, because it provides easy access for addition of components such as glucose during continuous kinetic measurements. Our current lab protocols for measurement of vacuolar and cytosolic pH are listed below; both are also easily adapted to microplate assays.

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments (optional)</td>
 </tr>
 <tr>
 <td>Spectrofluorometer</td>
 <td>Horiba Jobin Yvon</td>
 <td>Model Fluoromax-4</td>
 <td>Temperature control and stirring capability are desirable.</td>
 </tr>
 <tr>
 <td>BCECF-AM</td>
 <td>Invitrogen/Molecular Probes</td>
 <td>B1150</td>
 <td>Prepare a 12 mM stock in dry DMSO, store as aliquots at -20 &deg;C </td>
 </tr>
 <tr>
 <td>monensin</td>
 <td>Sigma</td>
 <td>M5273</td>
 <td>Toxic. </td>
 </tr>
 <tr>
 <td>nigericin</td>
 <td>Sigma</td>
 <td>N7143</td>
 <td>Toxic. </td>
 </tr>
 <tr>
 <td>MES</td>
 <td>Sigma</td>
 <td>M8250</td>
 <td></td>
 </tr>
 </tbody>
</table>

measurement of vacuolar and cytosolic ph in vivo in yeast cell suspensions

Vacuolar and cytosolic pH are highly regulated in yeast cells and occupy a central role in overall pH homeostasis. We describe protocols for ratiometric measurement of pH in vivo using pH-sensitive fluorophores localized to the vacuole or cytosol. Vacuolar pH is measured using BCECF, which localizes to the vacuole in yeast when introduced into cells in its acetoxymethyl ester form. Cytosolic pH is measured with a pH-sensitive GFP expressed under control of a yeast promoter, yeast pHluorin. Methods for measurement of fluorescence ratios in yeast cell suspensions in a fluorimeter are described. Through these protocols, single time point measurements of pH under different conditions or in different yeast mutants have been compared and changes in pH over time have been monitored. These methods have also been adapted to a fluorescence plate reader format for high-throughput experiments. Advantages of ratiometric pH measurements over other approaches currently in use, potential experimental problems and solutions, and prospects for future use of these techniques are also described.

Figure 1 presents vacuolar pH data obtained on wild-type yeast cells grown in rich medium (yeast extract-peptone-dextran; YEPD) buffered to pH 5 with 50 mM MES. We often grow the cells in buffered medium because the pH of the medium can change quite dramatically during overnight growth, particularly for minimal medium, and we have found that the pH of the growth medium can affect vacuolar pH responses 3. However, it is also acceptable for many experiments to grow the cells in unbuffered...

Watch this Scientific Journal Video about Measurement of Vacuolar and Cytosolic pH In Vivo in Yeast Cell Suspensions at JoVE.com

Measurement of Vacuolar and Cytosolic pH In Vivo in Yeast Cell Suspensions

Vacuolar and cytosolic pH can be measured in live yeast (S. cerevisiae) cells using ratiometric fluorescent dyes localized to specific cellular compartments. We describe procedures for measuring vacuolar pH with BCECF-AM, which localizes to the vacuole in yeast, and cytosolic pH with a cytosolic ratiometric pH-sensitive GFP (yeast pHluorin).

Measurement of Vacuolar and Cytosolic pH In Vivo in Yeast Cell Suspensions

Vacuolar and  cytosolic pH are highly regulated in yeast cells and occupy a central role in  overall pH homeostasis. We describe  protocols for ...

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19.8K Views.  SUNY Upstate Medical University. Vacuolar and cytosolic pH can be measured in live yeast (S. cerevisiae) cells using ratiometric fluorescent dyes localized to specific cellular compartments. We describe procedures for measuring vacuolar pH with BCECF-AM, which localizes to the vacuole in yeast, and cytosolic pH with a cytosolic ratiometric pH-sensitive GFP (yeast pHluorin).

Video: Measurement of Vacuolar and Cytosolic pH In Vivo in Yeast Cell Suspensions

In vivo Micro-circulation Measurement in Skeletal Muscle by Intra-vital Microscopy

BACKGROUND: Regulatory factors and detailed physiology of in vivo microcirculation have remained not fully clarified after many different modalities of imaging had invented. While many macroscopic parameters of blood flow reflect flow velocity, changes in blood flow velocity and red blood cell (RBC) flux does not hold linear relationship in the microscopic observations. There are reports of discrepancy between RBC velocity and RBC flux, RBC flux and plasma flow volume, and of spatial and temporal heterogeneity of flow regulation in the peripheral tissues in microscopic observations, a scientific basis for the requirement of more detailed studies in microcirculatory regulation using intravital microscopy. METHODS: We modified Jeff Lichtman's method of in vivo microscopic observation of mouse sternomastoid muscles. Mice are anesthetized,
ventilated, and injected with PKH26L-fluorescently labeled RBCs for microscopic observation.RESULT &amp; CONCLUSIONS: Fluorescently labeled RBCs are detected and distinguished well by a wide-field microscope. Muscle contraction evoked by electrical stimulation induced increase in RBC flux. Quantification of other parameters including RBC velocity and capillary density were feasible. Mice tolerated well the surgery, injection of stained RBCs, microscopic observation, and electrical stimulation. No muscle or blood vessel damage was observed, suggesting that our method is relatively less invasive and suited for long-term observations.

A new versatile method for observation of microcirculation is presented. It is considered suitable for long-term observation, and for combination with pharmacophysiological or molecular biological interventions.

BACKGROUND: Regulatory factors and detailed physiology of in vivo microcirculation have remained not fully clarified after many different modalities of ...

Single-cell Suction Recordings from Mouse Cone Photoreceptors

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment are open and allow cations to flow steadily inwards across the membrane, depolarizing the cell. Light exposure triggers the closure of the CNG channels, blocks the inward cation current flow, and thus results in cell hyperpolarization. Based on the polarity of photoreceptors, a suction recording method was developed in 1970s that, unlike the classic patch-clamp technique, does not require penetrating the plasma membrane 1. Drawing the outer segment into a tightly-fitting glass pipette filled with extracellular solution allows recording the current changes in individual cells upon test-flash exposure. However, this well-established "outer-segment-in (OS-in)" suction recording is not suitable for mouse cone recordings, because of the low percentage of cones in the mouse retina (3%) and the difficulties in identifying the cone outer segments. Recently, an inner-segment-in (IS-in) recording configuration was developed to draw the inner segment/nuclear region of the photoreceptor into the recording pipette 2,3. In this video, we will show how to record from individual mouse cone photoresponses using single-cell suction electrode.

We will show how to record flash responses from single mouse cones using a suction electrode.

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment ...

Quantitative Measurement of Invadopodia-mediated Extracellular Matrix Proteolysis in Single and Multicellular Contexts

Cellular invasion into local tissues is a process important in development and homeostasis. Malregulated invasion and subsequent cell movement is characteristic of multiple pathological processes, including inflammation, cardiovascular disease and tumor cell metastasis1. Focalized proteolytic degradation of extracellular matrix (ECM) components in the epithelial or endothelial basement membrane is a critical step in initiating cellular invasion. In tumor cells, extensive in vitro analysis has determined that ECM degradation is accomplished by ventral actin-rich membrane protrusive structures termed invadopodia2,3. Invadopodia form in close apposition to the ECM, where they moderate ECM breakdown through the action of matrix metalloproteinases (MMPs). The ability of tumor cells to form invadopodia directly correlates with the ability to invade into local stroma and associated vascular components3. 
Visualization of invadopodia-mediated ECM degradation of cells by fluorescent microscopy using dye-labeled matrix proteins coated onto glass coverslips has emerged as the most prevalent technique for evaluating the degree of matrix proteolysis and cellular invasive potential4,5. Here we describe a version of the standard method for generating fluorescently-labeled glass coverslips utilizing a commercially available Oregon Green-488 gelatin conjugate. This method is easily scaled to rapidly produce large numbers of coated coverslips. We show some of the common microscopic artifacts that are often encountered during this procedure and how these can be avoided. Finally, we describe standardized methods using readily available computer software to allow quantification of labeled gelatin matrix degradation mediated by individual cells and by entire cellular populations. The described procedures provide the ability to accurately and reproducibly monitor invadopodia activity, and can also serve as a platform for evaluating the efficacy of modulating protein expression or testing of anti-invasive compounds on extracellular matrix degradation in single and multicellular settings.

We describe the prototypical method for producing microscope coverslips coated with fluorescent gelatin for visualizing invadopodia-mediated matrix degradation. Computational techniques using available software are presented for quantifying the resultant levels of matrix proteolysis by single cells within a mixed population and for multicellular groups encompassing entire microscopic fields.

Cellular invasion into local tissues is a process important in development and homeostasis.  Malregulated invasion and subsequent cell movement is ...

Simple and Efficient Technique for the Preparation of Testicular Cell Suspensions

Mammalian testes are very complex organs that contain over 30 different cell types, including somatic testicular cells and different stages of germline cells. This heterogeneity is an important drawback concerning the study of the bases of mammalian spermatogenesis, as pure or enriched cell populations in certain stages of sperm development are needed for most molecular analyses1.
 Various strategies such as Staput2,3, centrifugal elutriation1, and flow cytometry (FC)4,5 have been employed to obtain enriched or purified testicular cell populations in order to enable differential gene expression studies. 
It is required that cells are in suspension for most enrichment/ purification approaches. Ideally, the cell suspension will be representative of the original tissue, have a high proportion of viable cells and few multinucleates - which tend to form because of the syncytial nature of the seminiferous epithelium6,7 - and lack cell clumps1 . Previous reports had evidenced that testicular cell suspensions prepared by an exclusively mechanical method clumped more easily than trypsinized ones1 . On the other hand, enzymatic treatments with RNAses and/or disaggregating enzymes like trypsin and collagenase lead to specific macromolecules degradation, which is undesirable for certain downstream applications. The ideal process should be as short as possible and involve minimal manipulation, so as to achieve a good preservation of macromolecules of interest such as mRNAs. Current protocols for the preparation of cell suspensions from solid tissues are usually time-consuming, highly operator-dependent, and may selectively damage certain cell types1,8 . 
	The protocol presented here combines the advantages of a highly reproducible and extremely brief mechanical disaggregation with the absence of enzymatic treatment, leading to good quality cell suspensions that can be used for flow cytometric analysis and sorting4, and ulterior gene expression studies9 .

A novel protocol for the mechanical preparation of testicular cell suspensions from rodent material, avoiding enzymes and detergents, is described. The method is very simple, fast, reproducible, and renders good quality cell suspensions, which are suitable for flow sorting and RNA extraction.

Mammalian testes are very complex organs that contain over 30 different cell types, including somatic testicular cells and different stages of germline ...

Simultaneous pH Measurement in Endocytic and Cytosolic Compartments in Living Cells using Confocal Microscopy

Intracellular pH is tightly regulated and differences in pH between the cytoplasm and organelles have been reported1. Regulation of cellular pH is crucial for homeostatic control of physiological processes that include: protein, DNA and RNA synthesis, vesicular trafficking, cell growth and cell division. Alterations in cellular pH homeostasis can lead to detrimental functional changes and promote progression of various diseases2. Various methods are available for measuring intracellular pH but very few of these allow simultaneous measurement of pH in the cytoplasm and in organelles. Here, we describe in detail a rapid and accurate method for the simultaneous measurement of cytoplasmic and organellar pH by using confocal microscopy on living cells3. This goal is achieved with the use of two pH-sensing ratiometric dyes that possess selective cellular compartment partitioning. For instance, SNARF-1 is compartmentalized inside the cytoplasm whereas HPTS is compartmentalized inside endosomal/lysosomal organelles. Although HPTS is commonly used as a cytoplasmic pH indicator, this dye can specifically label vesicles along the endosomal-lysosomal pathway after being taken up by pinocytosis3,4. Using these pH-sensing probes, it is possible to simultaneously measure pH within the endocytic and cytoplasmic compartments. The optimal excitation wavelength of HPTS varies depending on the pH while for SNARF-1, it is the optimal emission wavelength that varies. Following loading with SNARF-1 and HPTS, cells are cultured in different pH-calibrated solutions to construct a pH standard curve for each probe. Cell imaging by confocal microscopy allows elimination of artifacts and background noise. Because of the spectral properties of HPTS, this probe is better suited for measurement of the mildly acidic endosomal compartment or to demonstrate alkalinization of the endosomal/lysosomal organelles. This method simplifies data analysis, improves accuracy of pH measurements and can be used to address fundamental questions related to pH modulation during cell responses to external challenges.

Several methods are available for measuring intracellular pH but very few of these allow simultaneous measurement of cytoplasmic and organellar pH. Here, we describe in detail a rapid and accurate methodology to simultaneously measure cytoplasmic and vesicular pH by ratiometric imaging of living cells.

Intracellular pH is tightly regulated and differences in pH between the cytoplasm and organelles have been reported1. Regulation of cellular pH is crucial ...

Measurement of Heme Synthesis Levels in Mammalian Cells

Heme serves as the prosthetic group for a wide variety of proteins known as hemoproteins, such as hemoglobin, myoglobin and cytochromes. It is involved in various molecular and cellular processes such as gene transcription, translation, cell differentiation and cell proliferation. The biosynthesis levels of heme vary across different tissues and cell types and is altered in diseased conditions such as anemia, neuropathy and cancer. This technique uses [4-14C] 5-aminolevulinic acid ([14C] 5-ALA), one of the early precursors in the heme biosynthesis pathway to measure the levels of heme synthesis in mammalian cells. This assay involves incubation of cells with [14C] 5-ALA followed by extraction of heme and measurement of the radioactivity incorporated into heme. This procedure is accurate and quick. This method measures the relative levels of heme biosynthesis rather than the total heme content. To demonstrate the use of this technique the levels of heme biosynthesis were measured in several mammalian cell lines.

Altered intracellular heme levels are associated with common diseases such as cancer. Thus, there is a need to measure heme biosynthesis levels in diverse cells. The goal of this protocol is to provide a fast and sensitive method to measure and compare the levels of heme synthesis in different cells.

Heme serves as the prosthetic group for a wide variety of proteins known as hemoproteins, such as hemoglobin, myoglobin and cytochromes. It is involved in ...

Analysis of Cell Suspensions Isolated from Solid Tissues by Spectral Flow Cytometry

Flow cytometry has been used for the past 40 years to define and analyze the phenotype of lymphoid and other hematopoietic cells. Initially restricted to the analysis of a few fluorochromes, currently there are dozens of different fluorescent dyes, and up to 14-18 different dyes can be combined at a time. However, several limitations still impair the analytical capabilities. Because of the multiplicity of fluorescent probes, data analysis has become increasingly complex due to the need of large, multi-parametric compensation matrices. Moreover, mutant mouse models carrying fluorescent proteins to detect and trace specific cell types in different tissues have become available, so the analysis (by flow cytometry) of auto-fluorescent cell suspensions obtained from solid organs is required. Spectral flow cytometry, which distinguishes the shapes of emission spectra along a wide range of continuous wavelengths, addresses some of these problems. The data is analyzed with an algorithm that replaces compensation matrices and treats auto-fluorescence as an independent parameter. Thus, spectral flow cytometry should be capable of discriminating fluorochromes with similar emission peaks and can provide a multi-parametric analysis without compensation requirements.
This protocol describes the spectral flow cytometry analysis, allowing for a 21-parameter (19 fluorescent probes) characterization and the management of an auto-fluorescent signal, providing high resolution in minor population detection. The results presented here show that spectral flow cytometry presents advantages in the analysis of cell populations from tissues difficult to characterize in conventional flow cytometry, such as the heart and the intestine. Spectral flow cytometry thus demonstrates the multi-parametric analytical capacity of high-performing conventional flow cytometry without the requirement for compensation and enables auto-fluorescence management.

This article describes spectral cytometry, a new approach in flow cytometry that uses the shapes of emission spectra to distinguish fluorochromes. An algorithm replaces compensations and can treat auto-fluorescence as an independent parameter. This new approach allows for the proper analysis of cells isolated from solid organs.

Flow cytometry has been used for the past 40 years to define and analyze the phenotype of lymphoid and other hematopoietic cells. Initially restricted to ...

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 ...

4D Microscopy of Yeast

The goal of this protocol is to characterize how membrane compartments form and transform in live cells of budding yeast. Many intracellular compartments in yeast are dynamic, and a full understanding of their properties requires time-lapse imaging. Multi-color 4D confocal fluorescence microscopy is a powerful method for tracking the behavior and composition of an intracellular compartment on a time scale of 5-15 min. Rigorous analysis of compartment dynamics requires the capture of thousands of optical sections. To achieve this aim, photobleaching and phototoxicity are minimized by scanning rapidly at very low laser power, and the pixel dimensions and Z-step intervals are set to the largest values that are compatible with sampling the image at full resolution. The resulting 4D data sets are noisy but can be smoothed by deconvolution. Even with high quality data, the analysis phase is challenging because intracellular structures are often numerous, heterogeneous, and mobile. To meet this need, custom ImageJ plugins were written to array 4D data on a computer screen, identify structures of interest, edit the data to isolate individual structures, quantify the fluorescence time courses, and make movies of the projected Z-stacks. 4D movies are particularly useful for distinguishing stable compartments from transient compartments that turn over by maturation. Such movies can also be used to characterize events such as compartment fusion, and to test the effects of specific mutations or other perturbations.

This protocol describes the analysis of fluorescently labeled intracellular compartments in budding yeast using multi-color 4D (time-lapse 3D) confocal microscopy. The imaging parameters are chosen to capture adequate signals while limiting photodamage. Custom ImageJ plugins allow labeled structures to be tracked and quantitatively analyzed.

The goal of this protocol is to characterize how membrane compartments form and transform in live cells of budding yeast. Many intracellular compartments ...

Effect of Fluorescent Proteins on Fusion Partners Using Polyglutamine Toxicity Assays in Yeast

For the investigation of protein localization and trafficking using live cell imaging, researchers often rely on fusing their protein of interest to a fluorescent reporter. The constantly evolving list of genetically encoded fluorescent proteins (FPs) presents users with several alternatives when it comes to fluorescent fusion design. Each FP has specific optical and biophysical properties that can affect the biochemical, cellular, and functional properties of the resulting fluorescent fusions. For instance, several FPs tend to form nonspecific oligomers that are susceptible to impede on the function of the fusion partner. Unfortunately, only a few methods exist to test the impact of FPs on the behavior of the fluorescent reporter. Here, we describe a simple method that enables the rapid assessment of the impact of FPs using polyglutamine (polyQ) toxicity assays in the budding yeast Saccharomyces cerevisiae. PolyQ-expanded huntingtin proteins are associated with the onset of Huntington&#39;s disease (HD), where the expanded huntingtin aggregates into toxic oligomers and inclusion bodies. The aggregation and toxicity of polyQ expansions in yeast are highly dependent on the sequences flanking the polyQ region, including the presence of fluorescent tags, thus providing an ideal experimental platform to study the impact of FPs on the behavior of their fusion partner.

This article describes protocols to assess the effect of fluorescent proteins on the aggregation and toxicity of misfolded polyglutamine expansion for the rapid evaluation of a newly uncharacterized fluorescent protein in the context of fluorescent reporters.

For the investigation of protein localization and trafficking using live cell imaging, researchers often rely on fusing their protein of interest to a ...

Measurement of Vacuolar and Cytosolic pH In Vivo in Yeast Cell Suspensions

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Chapters in this video

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Single-cell Suction Recordings from Mouse Cone Photoreceptors

Quantitative Measurement of Invadopodia-mediated Extracellular Matrix Proteolysis in Single and Multicellular Contexts

Simple and Efficient Technique for the Preparation of Testicular Cell Suspensions

Simultaneous pH Measurement in Endocytic and Cytosolic Compartments in Living Cells using Confocal Microscopy

Measurement of Heme Synthesis Levels in Mammalian Cells

Analysis of Cell Suspensions Isolated from Solid Tissues by Spectral Flow Cytometry

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

4D Microscopy of Yeast

Effect of Fluorescent Proteins on Fusion Partners Using Polyglutamine Toxicity Assays in Yeast