Current Protocols in Cytometry
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Functional analysis of cellular organelles can be accomplished by staining cells with suitable organelle-specific dyes and then analyzing the fluorescence of the stained cells with a flow cytometer. With this methodology it is possible to resolve suspected heterogeneity in organelle function or content within a population of cells. Flow cytometry does not provide morphological information; if that is desired, quantitative microscopy­using a video microscope with digital image analysis system, or a confocal microscope­should be employed. LYSOSOMES Lysosomes are organelles devoted to degradation of cellular macromolecules and subsequent recycling of their components. The degradative enzymes of lysosomes all operate at an acidic pH. These features offer two ways of specifically staining lysosomes. First, basic fluorophores that are plasma membrane permeant will accumulate inside lysosomes. That means that the amount of fluorescence obtained with a plasma membrane-permeant base will be related to the amount of lysosomal mass in a cell. Second, the enzymatic activity of lysosomes can be monitored using nonfluorescent dyes containing a chemical group that can be removed by an acidic hydrolase to generate a fluorophore. Procedures for assessing the amount of lysosomal mass in a cell using LysoTracker dyes, which are plasma membrane-permeant bases (see Basic Protocol 2), and for staining cells with a fluorogenic lysosomal enzyme substrate (see Alternate Protocol 2) are described. As staining is dependent upon lysomal activity, fixation is not possible. BASIC PROTOCOL 2 STAINING OF LIVE CELLS FOR MEASUREMENT OF LYSOSOMAL MASS OR FUNCTION BY FLOW CYTOMETRY OR MICROSCOPY This protocol presents two methods for staining cultured mammalian cells with fluorescent plasma membrane-permeant bases: one performed with cells in suspension, suitable for subsequent analysis by flow cytometry, and one performed with cells on coverslips, suitable for subsequent microscopic analysis. The protocols can be adapted to staining of plant and yeast cells. Samples can be analyzed with flow cytometers equipped with an excitation light source matching the dye excitation wavelength range (see Table 9.4.2) or with microscopes equipped with appropriate filters. This protocol requires basic expertise in cell handling, microscopy, and flow cytometry. Materials

Cells in suspension (APPENDIX 3B; for flow cytometry procedure) or grown in monolayer (APPENDIX 3B) on coverslips (for microscopy procedure)

Cell culture medium supplemented with 10% FBS (APPENDIX 2A), 37°C

100 ´ LysoTracker Blue, Green, Yellow, or Red dye working solution (see recipe)

15 mM propidium iodide (aqueous; stable >1 year at 4°C) or 50 mM SYTOX Green working solution (prepare immediately before use from purchased stock solution)

9:1 (v/v) HBSS/FBS, 37°C (freshly prepared, for microscopy; see APPENDIX 2A for HBSS recipe)

Melted wax or nail polish (for microscopy)

15-ml screw-cap centrifuge tubes and 12 ´ 75-mm polypropylene tubes (for flow cytometry)

18 ´ 18-mm coverslips sterilized by dipping into absolute ethanol and subsequent flaming (for microscopy)

35-mm cell culture dishes (for microscopy)

Flow cytometer with either a mercury arc lamp, an argon-ion laser, or a HeNe laser as excitation source, or fluorescence microscope (UNIT 2.4)

Computer for data collection and processing

Additional reagents and equipment for cell culture and harvesting (APPENDIX 3B)

1. a. Harvest cultured cells by standard procedures into 15-ml screw-cap centrifuge tubes and centrifuge 5 min at 200 ´g, room temperature.
    
2. a. Resuspend the cell pellet at 0.5-1.0 ´ 10⁶ cells/ml in 37°C cell culture medium. Leave cell suspension at 37°C for ³5 min.
    
   Because the functional state of the lysosomes is to be monitored, it is advisable to keep cell suspensions at their optimal temperature (37°C) and allow them to recover for a brief moment after harvesting.
    
3. a. Thaw out LysoTracker dye working solution at room temperature, keeping it protected from light (e.g., in a drawer).
    
   Dye solutions decompose rapidly if exposed to light. Do not refreeze thawed vials, because the dyes decompose during freeze-thaw cycles.
    
4. a. Divide cell suspension into 1-ml aliquots in 12 ´ 75-mm polypropylene tubes, add 10 ml dye working solution, and mix immediately by briefly vortexing at maximum speed.
    
   Dye concentrations in the range of 0.05 to 0.1 µM are recommended, because nonlysosomal staining may occur at higher concentrations.
    
5. a. Incubate 15 to 30 min at 37°C in the dark or in subdued light. After staining, place tube in a melting ice bath.
    
6. a. Optional: Exclude dead cells by costaining ³15 min at room temperature with 5 mM propidium iodide (final) if LysoTracker Blue, Green, or Yellow is used or 0.5 mM SYTOX Green (final) if LysoTracker Red is used.
    
7. a. Set up and optimize the flow cytometer (see Table 9.4.2). Because of the wide variation in cellular lysosome content, use logarithmic signal amplification for the signal channels collecting lysosome-related fluorescence.
    
   Use a bandpass filter centered around 620 nm for propidium iodide or around 530 nm for SYTOX Green.
    
8. a. Carefully resuspend the cell sample by gently pipetting up and down a few times immediately before analysis.
    
   Cells tend to clump during staining; to obtain meaningful data on a per-cell basis, it is essential to resuspend cells immediately before analysis.
    

1. b. Culture adherent cells overnight on sterilized 18 ´ 18-mm coverslips.
    
   Alternatively, larger coverslips and culture dishes can be used; in this case the volumes of the staining and other solutions should be adjusted accordingly. This protocol assumes that 35-mm dishes are used throughout.
    
2. b. Take coverslip with cells out of the cell culture dish, rinse once with 37°C cell culture medium, and place in a 35-mm cell culture dish with 1 ml of 37°C cell culture medium.
    
3. b. Thaw out LysoTracker dye working solution at room temperature, keeping it protected from light (e.g., in a drawer).
    
   Dye solutions decompose rapidly if exposed to light. Do not refreeze thawed-out vials because the dyes decompose during freeze-thaw cycles.
    
4. b. Add 10 ml dye working solution to the dish and swirl immediately to distribute the dye evenly.
    
5. b. Incubate 15 to 30 min at 37°C in the dark or in subdued light. After staining, briefly rinse three times with 37°C 9:1 (v/v) HBSS/FBS.
    
   At this stage, cells can be labeled with propidium iodide or SYTOX Green (see Basic Protocol 2, step 6a) to distinguish dead cells; however, these generally can be distinguished by morphology.
    
6. b. Invert coverslip and mount onto a slide in a drop of 9:1 HBSS/FBS while leaving some clearance. Do not apply pressure. Seal the coverslip by a method regularly used in the laboratory (e.g., melted wax or nail polish).
    
7. b. To observe cells stained with LysoTracker dyes by fluorescence microscopy, use excitation and emission bandpass filters covering wavelength ranges compatible with the data displayed in Table 9.4.2.
    
   Use a bandpass filter centered around 620 nm for propidium iodide or around 530 nm for SYTOX Green.
    

5- to 10-mM fluorescein di-b-D-galactopyranoside (FDG) working solution (in culture medium; prepared fresh from purchased stock solution)

1. Harvest cultured cells by standard procedures in 15-ml screw-cap centrifuge tubes and centrifuge 5 min at 200 ´g, room temperature.
    
2. Resuspend the cell pellet at 0.5-1.0 ´ 10⁶ cells/ml in 37°C cell culture medium. Leave cell suspensions at 37°C for ³5 min.
    
   Because the functional state of the lysosomes is to be monitored, it is advisable to keep cell suspensions at their optimal temperature (37°C) and to allow them to recover for a brief moment after harvesting.
    
3. Thaw out FDG working solution at room temperature, keeping it protected from light (e.g., in a drawer).
    
   Dye solutions decompose rapidly if exposed to light; FDG solutions have been found to decompose during repeated freeze-thaw cycles.
    
4. Divide cell suspension into 1-ml aliquots in 12 ´ 75-mm polypropylene tubes, add 10 ml FDG working solution, and mix immediately by briefly vortexing at maximum speed.
    
   Dye concentrations in the range of 50 to 100 µM are recommended to obtain a sufficiently strong signal. FDG working solution should be prepared immediately before use and kept on ice. It is stable for only 1 to 2 hours.
    
5. Incubate 15 to 30 min at 37°C in the dark or in subdued light. After staining, place tube in a melting ice bath.
    
6. Optional: Exclude dead cells by costaining ³15 min at room temperature with 5 mM propidium iodide (final).
    
7. Set up and optimize the flow cytometer (see Table 9.4.2). Because of the wide variation in lysosomal enzyme activity, use logarithmic signal amplification.
    
   Use a bandpass filter centered around 620 nm for detection of propidium iodide.
    
8. Carefully resuspend the cell sample by gently pipetting up and down a few times immediately before analysis.
    
   Cells tend to clump during staining; to obtain meaningful data on a per-cell basis, it is essential to resuspend cells immediately before analysis.
    

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