Thanks are due to the Tyrolean Science Fund (TWF) for providing grant #256524 to AL, to the Vienna Science and Technology Fund (WWTF) for providing grant #LS13-086 to SZ, and to the Publication Fund of the University of Innsbruck for supporting open access publication. The authors also thank the Department of Zoology of the University of Innsbruck for providing the Leica TCS SP5 II confocal laser scanning microscope.

1. Preparation of fungal samples
<ol>
	<li>Fungal pre-cultures
	<ol>
		<li>Inoculate the desired strain on an appropriate solid agar medium, such as potato dextrose agar (PDA) for Trichoderma atroviride or Vogel&#8217;s Minimal Medium (VMM) for Neurospora crassa. Add an appropriate selection marker when working with transformant strains.</li>
		<li>Incubate the pre-culture at the organism&#8217;s optimal temperature. For instance, T. atroviride at 25 &#176;C and N. crassa at 30 &#176;C, and 12 h/12 h light/dark cycles until a sporulating mycelium has developed but not yet reached the edge of the plate. On a standard size Petri dish (9.2 cm &#216;), this takes the wild-type T. atroviride on 4&#8211;6 days average, whereas the N. crassa wild type reaches this stage after 3&#8211;4 days on average.</li>
	</ol>
	</li>
	<li>Cultivation of fungal colonies
	<ol>
		<li>Using a sterile scalpel, cut a small 3 mm x 3 mm agar block carrying non-sporulating mycelium from the colony edge of the pre-culture.</li>
		<li>Place the agar block at the center of a fresh solid medium plate to inoculate the experimental culture.</li>
		<li>Incubate the experimental culture according to the developmental stage intended to be investigated. For instance, the wild-type T. atroviride requires 20&#8211;22 hours at 25 &#176;C in the dark to develop colonies of about 2 cm diameter on PDA, whereas the wild-type N. crassa reaches colony diameters of about 4 cm after 14&#8211;16 h of incubation at 30 &#176;C in the dark on VMM. 
		NOTE: Incubation in the dark prevents the formation of pigments which might introduce autofluorescence. In order to eliminate medium background fluorescence from the experimental culture, replace agar with 1.5% w/v of a transparent solidifying agent (see the Table of Materials), and any complex medium with a defined minimal medium.</li>
	</ol>
	</li>
	<li>Cultivation of solid germling cultures
	<ol>
		<li>Use 5 mL of sterile physiological salt solution (0.9% w/v NaCl) to harvest conidial spores from the pre-culture plate and collect the resulting spore suspension in a 15 mL screw cap tube.</li>
		<li>Mix the spore suspension well by vigorous vortexing and subsequently filter it over a 1 cm x 5 cm strip of sterile filter fabric (see the Table of Materials) lightly stuffed into a 1 mL pipette tip (both assembled and autoclaved beforehand) into a fresh sterile tube.</li>
		<li>Determine the spore density with a cell counting chamber and prepare a 1 x 107 cells/mL spore suspension with physiological salt solution. 
		NOTE: The spore suspension can be kept at 4 &#176;C for up to two weeks.</li>
		<li>Prepare a standard size Petri dish (9.2 cm &#216;) with 20 mL of solid medium and add 15&#8211;20 sterile glass beads (3 mm &#216;) on top.</li>
		<li>Pipette 200 &#181;L of the spore suspension onto the medium plate and evenly distribute the cells across the whole plate by gentle shaking. Collect the glass beads into a beaker with 70% ethanol for reuse.</li>
		<li>Incubate the experimental culture according to the developmental stage intended to be investigated. For instance, the T. atroviride wild type requires 5&#8211;6 h at 25 &#176;C in the dark to develop conidial germlings on PDA, whereas the N. crassa wild type develops conidial germlings after 3&#8211;4 h of incubation at 30 &#176;C in the dark on VMM. 
		NOTE: In order to eliminate any medium background fluorescence from the experimental culture, replace agar with 1.5% w/v of a transparent solidifying agent, and any complex medium with a defined minimal medium.</li>
	</ol>
	</li>
	<li>Cultivation of liquid germling cultures
	<ol>
		<li>Fill 190 &#181;L of liquid culture medium into each well of an 8-well chambered micro-slide.</li>
		<li>Add 10 &#181;L of a 1 x 107 cells/mL spore solution (prepared in steps 1.4.1&#8211;1.4.3) and mix by gently pipetting up-and-down a few times. The resulting total number of cells is 1 x 105 per well.</li>
		<li>Incubate the experimental culture according to the developmental stage intended to be investigated. For instance, the wild type T. atroviride requires 5&#8211;6 h at 25 &#176;C in the dark to develop conidial germlings in potato dextrose broth (PDB), whereas the wild type N. crassa develops conidial germlings after 3&#8211;4 h of incubation at 30 &#176;C in the dark in liquid VMM.</li>
	</ol>
	</li>
</ol>
2. Preparation of dye working solutions
<ol>
	<li>To guarantee full solubility of each dye, prepare 2 mM stock solutions in dimethyl sulfoxide (DMSO) by adding the appropriate amount (see exact weights in Table 1) to 1 mL of 100% DMSO and mix well by vortexing. 
	CAUTION: Make sure to take the DMSO from a septum-sealed bottle; it should be a clear transparent liquid. Upon contact with air, DMSO turns brown&#8212;probably due to the oxidation of trace impurities&#8212;and might negatively affect cell growth or dye staining.</li>
	<li>Filter sterilize the stock solution through a 0.2 &#181;m syringe membrane filter into a fresh sterile 1.5 mL reaction tube. To minimize dye bleaching, wrap the tube in aluminum foil. 
	NOTE: The dye stock solution can be aliquoted into smaller volumes to avoid thawing/freezing cycles, and kept at 4 &#176;C for several months.</li>
	<li>Prepare a 20 &#181;M aqueous dye working solution by dissolving 2 &#181;L of dye stock solution in 198 &#181;L of sterile distilled water in a fresh sterile 1.5 mL reaction tube. To minimize dye bleaching, wrap the tube in aluminum foil. 
	NOTE: The dye working solution should be prepared freshly on the day of the experiment.</li>
	<li>During sample mounting (see section 3), the dye working solution will be standardly diluted 1:10, resulting in a final dye concentration of 2 &#181;M and 0.1% w/v final DMSO concentration. 
	NOTE: Choosing different dilution factors by simply changing the volume ratio between dye working solution and mounting liquid, allows to easily adapt the desired final dye concentration. 
	CAUTION: To prevent unwanted effects due to dye or DMSO toxicity, the dilution factor should not fall under 1:4 to result maximum final concentrations of 5 &#181;M dye and 0.4% w/v DMSO. Higher dye concentrations will quickly saturate the system and prevent reliable signal quantification, whereas more than 0.5% w/v (&#8805; 62.5 mM) DMSO can impair cell development38.</li>
</ol>
3. Sample preparation for microscopy
<ol>
	<li>Mount samples from fungal colonies (step 1.2) or solid germling cultures (step 1.3) by the inverted agar block method.
	<ol>
		<li>Keep a clean 24 mm x 60 mm glass cover slip (#1 = 0.13&#8211;0.16 mm thickness) ready and add 18 &#181;L of liquid minimal medium (VMM or M9) or physiological salt solution onto the center.</li>
		<li>Add 2 &#181;L of the 20 &#181;M dye working solution to the 18 &#181;L of liquid and mix well by pipetting up and down several times, while avoiding the production of air bubbles. 
		NOTE: When working with several samples, it is advisable to prepare a master mix of liquid-dye solution for all in order to ensure equal dye concentration throughout the experiment.</li>
		<li>With a clean scalpel, cut out a 15 mm x 15 mm sample from the periphery of the colony or solid germling culture and place it vertically beside the medium drop onto the cover slip.</li>
		<li>Using the scalpel to support the upper edge of the block and a finger to hold the rear side of the block in place, slowly lower the side carrying the mycelium or germlings onto the liquid. The sample is now ready for transfer onto the microscope stage. 
		CAUTION: It is essential to do this slowly and very carefully in order to minimize mechanical stress on the cells and to avoid air bubbles being trapped between sample and cover slip.</li>
	</ol>
	</li>
	<li>Mount the liquid germling cultures from step 1.4. 
	NOTE: Most conveniently, liquid germling cultures in chambered micro-well slides can be directly transferred and further manipulated on the microscope stage.
	<ol>
		<li>Add 22 &#181;L of dye working solution to 200 &#181;L of liquid medium to result in standard final concentrations of 2 &#181;M dye and 0.1% w/v DMSO. 
		NOTE: Liquid germling cultures have the great advantage that fluorescent dyes (or other chemicals, such as inhibitors) can be added at any desired time point of the experiment, also during recording. In that case, special care needs to be taken to administer the liquid drops very slowly in order to not disturb the cells. System vibrations and Brownian motion might already introduce some cell movement.</li>
	</ol>
	</li>
</ol>
4. Live-cell microscopy
<ol>
	<li>Adjust the basic image acquisition settings. The following image acquisition settings allow to capture staining dynamics in individual hyphae and are applicable to both of the following assays
	<ol>
		<li>Apply 5&#8211;10% laser power of 20% of the full output power of the device.</li>
		<li>Use a Plan Apo 60x&#8211;63x glycerol or water immersion objective with a high numerical aperture &#8805; 1.2.</li>
		<li>Restrict the image acquisition area to the outline of the hyphae by setting an image size of 1024 x 256 pixels and by using an optical zoom factor of 2&#8211;3.</li>
		<li>Use bidirectional scanning with 400 Hz. Adjust the pinhole size to 1 Airy unit.</li>
		<li>Set the gain of the most sensitive detector to 100%.</li>
		<li>For time laps recording, start image acquisition with one frame every 15 s to allow reasonable temporal resolution without producing dye bleaching or photo stress.</li>
		<li>For 3D recording, set the upper and lower spatial limit to the boundary of the hyphae and space optical sections 1 &#181;m apart to allow reasonable spatial resolution. 
		NOTE: Due to the fast growth of hyphae, high spatial resolution in the Z axis has often been sacrificed for high temporal resolution in the X/Y axis or the other way around. Only very modern confocal laser scanning microscopes are fast enough to satisfy both demands.</li>
	</ol>
	</li>
	<li>Endocytosis uptake assays
	<ol>
		<li>Consult Figure 1 and Table 1 to identify the best excitation/emission settings for FM 1-43 and/or FM 4-64 available on the microscopy system and adjust accordingly. 
		NOTE: With the recommended 2 &#181;M concentration, incorporation of FM dye into the plasma membrane is instant in normal healthy cells. The whole process from initial plasma membrane staining to dye appearance in tubular vacuoles is usually completed within 30&#8211;45 min at room temperature. Increasing the FM dye concentration increases the S/N-ratio and thus produces higher contrast images quicker. However, it also speeds up the labeling process making it more difficult to differentiate the chronological succession of organelle staining.</li>
		<li>Start image recording using the above recommended basic image acquisition settings and evaluate the results.</li>
		<li>Optimize image acquisition settings to the spatial and temporal resolution required to capture the aspect of plasma membrane or endocytosis dynamics the experiment is focused on.</li>
		<li>For instance, in order to capture very fast dynamics in X/Y, decrease the overall image size, image only one focal plane and increase the scanning rate to 1 fps. For higher resolution in the Z axis, decrease resolution in X/Y, decrease image size and decrease the distance between optical sections to 0.5 &#181;m.</li>
	</ol>
	</li>
	<li>Cell wall dynamics
	<ol>
		<li>Consult Figure 4 and Table 1 to identify the best excitation/emission settings for the applied cell wall dye available on the microscopy system and adjust accordingly. 
		NOTE: Due to their broad emission spectra, CFW and SPF are not well suited for simultaneous co-imaging with other fluorophores, predominantly GFP. Some restrictions do even apply for sequential imaging approaches with these dyes, and thus have to be optimized individually.</li>
		<li>Start image recording using the above recommended basic image acquisition settings and evaluate the results. 
		NOTE: With the recommended 2 &#181;M concentration, incorporation of dye into the cell wall is not necessarily instant but reasonably fast. The whole process of septum formation, for instance, takes on average about 5&#8211;7 min at room temperature20. Increasing the cell wall dye concentration increases the S/N-ratio and thus produces higher contrast images quicker. However, it also rapidly introduces artefacts due to induced cell wall damage repair.</li>
		<li>Optimize image acquisition settings to the spatial and temporal resolution required to capture the aspect of cell wall morphogenesis the experiment is focused on, as outlined in section 4.2.3.</li>
	</ol>
	</li>
</ol>

The authors declare that they have no competing financial interests and nothing to disclose.

This article continues the ground breaking work that established the use of various fluorescent dyes as vital organelle markers for filamentous fungi in the early 2000s2,4,43, and attempts to discuss the photophysical and cell biological properties of FM dyes and selected cell wall dyes in greater detail than before. Especially with respect to unwanted cellular effects, such as membrane saturation or cell wall damage, that occur above certain dye concentrations. What previously has been considered non-toxic on the cellular level is now regarded toxic on the molecular level. Even though these effects might be very subtle and not directly evident by obvious changes in organelle or cell behavior, any possible interference of dye application other than visualization has to be minimized for the investigation of native molecular function. Fortunately, improved sensitivity and quantum efficiency of modern detectors, such as silicon avalanche photodiode detectors (Si-APDs)44 or the Airyscan area detector45, facilitate the use of even lower dye amounts than before. Another key objective of the article is to exemplify the co-imaging properties of these dyes with other fluorophores, most importantly, those of GFP as the most frequently used fluorescent protein in biology. This should aid the design of imaging experiments that aim to correlate the subcellular localization dynamics of fluorescent fusion proteins to those of the fungal cell wall, the plasma membrane or the endo- and exocytosis pathway etc.
Imaging under natural and stress-free conditions is key for the acquisition of reliable data. Some practical considerations regarding culture medium and sample preparation aim at providing a starting point for finding the optimal conditions that allow artefact-free, long time observation of healthy, unstressed cells with the highest S/N-ratio possible for any given sample. There is no universal way of achieving reliable and meaningful imaging results. It is inherent to the approach that biological variation of the sample, subjectivity and expectations of the microscopist, as well as image post processing have significant influence on data acquisition and interpretation, respectively. Hence, the practical experience of the microscopist, her/his intimate knowledge about the cell biology of the fungus under investigation, as well as skillful sample preparation to create conditions as &#8216;natural&#8217; and undisturbed as possible in a lab setting, are paramount for acquiring and evaluating imaging data that truthfully reflects the studied cellular phenomena. As a rule of thumb, the occurrence of unwanted side effects of fluorescent dyes, ranging from the subtle and thus not obviously visible activation of plasma membrane or cell wall re-modelling stress response pathways to the straightforward cytotoxic induction of cell autolysis, can only be securely prevented by applying low dye concentrations &#8804;2 &#181;M.
The application of fluorescent dyes is simple, yet their specificities are poorly characterized. A key strength of using fluorescent dyes is the preparative simplicity of the experimental protocols. Cultivation and sampling of the fungus, addition of the dye(s), and mounting onto the microscope stage are (with practice) straightforward. Adjustment of the basic imaging settings, including excitation and emission wavelengths, exposure times, time course settings etc., follow simple biophysical rules of the microscope and biological rules of the used fluorescent dyes inside the cells. Table 1 intends to support the identification of the most suitable dye or dye combination for experimentation. Furthermore, fluorescent dyes are reasonably priced, readily available with dependable high quality and thus ensure highly reproducible application.
Two major restrictions of using membrane or cell wall selective fluorescent dyes are the (often) limited knowledge of their precise staining properties, which in most cases are non-specific on the organelle and molecular level, and their concentration-dependent unwanted side effects. FM dyes are specific for lipid bilayers participating in endo- and exocytosis. However, precisely which subcellular organelles become successively labeled under the tested conditions is not immediately evident and requires comparison of different FM dye variants, and co-labeling with additional organelle-specific markers. The preference of FM 1-43 for mitochondrial membranes, in comparison to FM 4-64, is one example. Cell wall selective dyes display varying specificity for the three major polymers of the fungal cell wall. CFW is thought to be a non-specific stain for &#946;-glucans and chitin, SPF is thought to be most selective for &#946;-1,4-glucans, and CR is thought to be highly selective for &#945;- and &#946;-chitins. Information on the binding specificity of PFS to fungal cell wall polysaccharides is currently not available. To what ratio which fungal cell wall polymer is most effectively labeled at a given dye concentration in the fungal species under investigation is not easily answered, and the application of detailed measurements acquired in vitro or in vivo in other organisms or other fungal species must be considered very carefully. Unfortunately, this information is sparse and highly scattered in the literature35,42,46. More recent records that would follow on previous studies33 to provide new insights into the precise staining properties of the featured dyes specifically in fungi are currently not available.
Imaging controls are essential to evaluate staining patterns and cellular responses accurately. Probably the most challenging part, however, is to know the cell biology of the fungus so well that the recorded changes in the subcellular localization of membrane and cell wall selective fluorescent dyes, alterations in cellular architecture or hyphal growth pattern can exclusively and confidently be related to the intended effects of the experimental treatment. For this, it is crucial to have good controls alongside any new live-cell imaging experiment. These include the untreated wild type as negative imaging control to exclude background autofluorescence and detector noise from the acquired image, and to have a morphological comparator when working with mutants. Furthermore, a positive imaging control, for example a strain that expresses GFP or RFP in the cytoplasm or another well-known fluorescent marker protein, is essential to set the excitation light intensity to the required minimum and have a cell vitality control. Once these controls are set, the use of fluorescent dyes is not restricted to visualization tasks only, but their concentration-dependent staining dynamics, as well as concentration-dependent side effects can be analytically exploited; for instance, for quantitatively monitoring cell wall biosynthesis in real time or for the identification of mutant-specific phenotypes in susceptibility assays47.
Improved future applications depend on a detailed functional analyses of dye staining properties. A major ongoing challenge is to further improve and automate quantitative image analyses in order to advance functional evaluation of the subcellular dynamics of membrane and cell wall selective fluorescent dyes in filamentous fungi. For this, extensive, quantitative co-localization studies of these dyes with known organelle and cell wall polymer markers in combination with mutant strains deficient in particular transport pathways or that lack specific structural components are first required. Several endocytosis markers for comparative analyses with FM dyes are available48,49, and with respect to the still poorly characterized binding specificities of cell wall dyes in fungi, the application of fluorescently-labeled glucan-specific antibodies50 might provide one possibility to address this issue.

Twenty years ago, the way in which hyphal morphogenesis and the underlying molecular cell biology could be visualized in filamentous fungi was revolutionized by the application of the membrane selective fluorescent Fei Mao dye FM 4-641. Later, the benefit of the chitin-binding dye Calcofluor White as vital fluorescent marker of fungal cell wall dynamics was realized2. Since then, both dyes and variants thereof have become an inherent part of live-cell imaging analyses of organelle dynamics in fungi, and continue to provide unprecedented insights into the filamentous fungal lifestyle. This paper details the application of established and lesser-known fluorescent dyes for tracking plasma membrane dynamics, endo- and exocytosis and cell wall morphogenesis in filamentous fungi. Endocytosis tracking assays allow various cell biological questions related to the general study of endocytosis to be addressed3. For this, the localization, speed and succession of stained compartments upon FM dye addition is recorded by time lapse microscopy and quantitatively compared between the tested fungal strains4. Cell wall dyes delineate the outer boundary of the cell and allow tracking of morphogenetic events, including polarized hyphal tip growth2, hyphal branching5, hyphal fusion6,7 and septum formation8. Furthermore, they facilitate the quantification of localized cell wall deposition and the identification of defects during cell wall biogenesis9. Because detailed knowledge of the biochemical and biophysical properties of any fluorescent marker is a fundamental prerequisite for its successful in vivo application, these characteristics are first summarized for the six dyes featured in this article.
Membrane selective dyes 
FM (Fei Mao) styryl dyes are small amphiphilic molecules that cannot pass through but reversibly associate with the outer leaflet of the lipid bilayer of biological membranes10. They are virtually non-fluorescent in aqueous solution, but become intensely fluorescent upon plasma membrane integration, generating excellent signal-to-noise (S/N)-ratios11. These properties make them ideally suited for visualizing plasma membrane and intracellular organelle dynamics, including tracking of endo- and exocytosis12. The green-fluorescent FM 1-43 and the red-fluorescent FM 4-64 are the two most widely used fluorescent membrane markers for these purposes. SynaptoGreen C4 and SynaptoRed C2 are generic molecules from alternative suppliers that can be interchangeably used instead of FM 1-43 and FM 4-64, respectively.
Styryl dyes comprise three key structural regions: (1) the lipophilic tail that facilitates insertion of the dye into the lipid bilayer, (2) the fluorophore core that determines the spectral properties of the dye and is constituted by two aromatic rings connected by one to three double bonds, and (3) the positively charged hydrophilic head that prevents complete insertion and permeation of the dye through the membrane (Figure 1A).
The longer the lipophilic tail, the higher is the dye&#8217;s hydrophobicity and thus binding affinity to the membrane, but the lower is its water solubility and membrane de-staining rate. Consequently, different FM dye variants produce different staining dynamics and patterns. The higher hydrophobicity of the C4-tailed FM 1-43 provides a stronger and more stable fluorescence signal at plasma membranes and internal organelles quicker than the shorter C2-tailed FM 4-64, when applied at equimolar concentrations (Figure 2).
Importantly, the constant and high association/dissociation rates of both FM dyes11 with average retention times of 1&#8211;6 s per individual dye molecule13 reduce the chances for localized disruption of membrane function, for example, through the modification of membrane fluidity or forced permanent interaction of membrane proteins. This is probably the key reason why these molecules can be used as vital dyes. Nevertheless, FM dye concentrations above 50 &#181;M are toxic to fungal and plant cells2,14, and evidence from BY-2 tobacco protoplasts indicates that more than 20 &#181;M FM dye lead to plasma membrane saturation14. Thus, it is advisable not to exceed this limit, especially given the fact that excellent imaging has been achieved with as little as 2&#8211;5 &#181;M15,16.
Notably, the spectral properties of FM dyes vary greatly depending on the particular membrane microenvironment (reviewed14). Generally, excitation and emission spectra of FM dyes in pure solvent solutions (as usually provided in the product information) differ significantly from that in cellular environments and can, in most cases, not directly be consulted for selecting live-cell imaging settings. The excitation/emission maxima of FM 1-43 and FM 4-64, for instance, become blue-shifted by 37/46 nm and 43/64 nm, respectively, when bound to fungal membranes in comparison to their solutions in methanol (Table 1).
The ground-breaking fundamentals for the use of FM 4-64 and FM 1-43 for tracking plasma membrane, endo-/exocytosis and organelle dynamics, including the Spitzenk&#246;rper and mitochondria, have already been comprehensively documented for a wide range of filamentous fungal species previously2,4,17,18,19. Recommended imaging settings for both FM dyes that work in various filamentous fungal species are depicted in Figure 1B. Technical limitations of the available equipment or particular cellular and experimental conditions, such as culture medium, pH or temperature, however, might require some adaptations. Fortunately, FM dyes operate over a wide spectral range, and very good imaging results are achieved by exciting FM 1-43 with 514 nm or FM 4-64 with 488 nm. Consequently, the optimal imaging settings must be determined individually for each sample type and intended application.
The considerable Stoke&#8217;s shift of more than 135 nm of FM 4-64 allows excellent, simultaneous co-imaging with fluorophores emitting green light; this is frequently exploited to evaluate the intracellular localization dynamics of green fluorescent protein (GFP)-labeled fusion proteins relative to the plasma membrane and the endocytic pathway9,20.
Cell wall-selective dyes 
Calcofluor White M2R (CFW), also marketed as Fluorescent Brightener 28, is probably the best-known fluorescent dye used to stain the cell walls of bacteria, fungi, algae, higher plants and insects. Initially used as optical whitening agent in the paper, textile and detergent industry, its benefits for the clinical diagnosis of fungal infections was realized early on21,22. Because CFW intercalates irreversibly into the nascent chitin chain it disturbs normal chitin microfibril assembly during cell wall biogenesis thereby generating cell wall stress23. This in turn triggers a cell wall damage repair mechanism leading to locally heightened cell wall deposition as the result of glucan and chitin synthase activation24,25. This phenomenon can occur with any dye that operates by stably binding to cell wall polymers, is concentration-dependent and is most noticeable at hyphal tips which represent the most prolific growing and thus most sensitive parts of the mycelium (Figure 3). A comprehensive summary of the molecular machinery that responds to cell wall damage has recently been provided26.
Overdosed dye in combination with phototoxicity can lead to rapid cell lysis of hyphal compartments (Movie 1). Nonetheless, increased sensitivity to dye concentrations that are &#8220;vital&#8221; in the wild type can be exploited to identify defects in cell wall biosynthesis of gene loss-of-function mutants9. For CFW and Congo Red (CR), another textile colorant also known as Direct Red 28 and employed as &#945;- and &#946;-chitin-specific cell wall stain for fungi and insects27,28, threshold concentrations that strongly induce chitin synthases have been determined with &#62; 60 &#181;M CFW and &#62; 70 &#181;M CR, respectively, whereas concentrations &#60;15 &#181;M of either dye did not alter or inhibit fungal growth29,30,31. Hickey et al. placed this threshold concentration for CFW at 25 &#181;M2. Therefore, it is advisable to use dye concentrations &#8804; 5 &#181;M to exclude stress-related artefacts and ensure using these molecules as truly &#8220;vital fluorescent dyes&#8221;2,32. This equally applies to Solophenyl Flavine 7GFE 500 (SPF) and Pontamine Fast Scarlet 4B (PFS), synonymous to Direct Yellow 86 and Direct Red 23, respectively, two other useful cell wall dyes whose application for fungi has been reported for the first time more than a decade ago33. But despite their remarkable spectral properties34,35, the use of both dyes has since then been very limited36,37. As previously shown for 1.5 &#181;M CFW2, 2 &#181;M SPF are sufficient to resolve cell wall dynamics under native conditions with very high temporal resolution (Movie 2). The same results can be obtained with 2 &#181;M CR or PFS.
Together, these four dyes, CFW, SPF, PFS and CR, comprise a set of cell wall-selective fluorescent markers that cover almost the complete visible emission light spectrum (400&#8211;700 nm) utilized on modern fluorescent microscopes (Figure 4). The significant increase in fluorescence intensity upon binding to cell wall polymers is inherent to all four and generates excellent S/N-ratios. This in turn permits to keep dye concentrations and excitation light intensity very low and allows to perform cell wall staining as &#8220;low dose&#8221; live-cell imaging technique2. Because these cell wall dyes are plasma membrane impermeable, they simultaneously function as live/dead stains. Notably, due to their extremely wide emission light spectra, some limitations regarding the co-imaging properties of CFW and SPF with other fluorophores need to be carefully considered.

<table><tbody><tr><td>BRAND cell counting chamber</td><td>Merck</td><td>BR718005</td><td>Thoma format</td></tr><tr><td>Calcofluor White M2R</td><td>Merck/Sigma-Aldrich</td><td>F3543</td><td>cell wall dye</td></tr><tr><td>CFI Plan Apo VC 60x/1.2 NA XC WI</td><td>Nikon</td><td>MRD07602</td><td>water immersion objective</td></tr><tr><td>CFI Plan Apo VC 60x/1.2 NA XC WI</td><td>Nikon</td><td>MRD07602</td><td>water immersion objective</td></tr><tr><td>Congo Red</td><td>Merck/Sigma-Aldrich</td><td>C6277</td><td>cell wall dye</td></tr><tr><td>Dimethyl sulfoxide</td><td>VWR</td><td>8,36,73,230</td><td>organic solvent</td></tr><tr><td>Eclipse TE2000-E with C1 scanning unit</td><td>Nikon</td><td>custom configuration</td><td>inverted laser scanning confocal microscope used to acquire example data 2 and 4</td></tr><tr><td>Eclipse TE2000-U with Bio-Rad Radiance 2100 scannig unit</td><td>Nikon</td><td>custom configuration</td><td>inverted laser scanning confocal microscope used to acquire example data 5</td></tr><tr><td>FM 1-43</td><td>Merck/Sigma-Aldrich</td><td>S6814</td><td>membrane dye</td></tr><tr><td>FM 4-64</td><td>Merck/Sigma-Aldrich</td><td>S6689</td><td>membrane dye</td></tr><tr><td>Glass beads</td><td>Rettberg</td><td>1340691030</td><td>3 mm glass beads</td></tr><tr><td>Glass cover slips</td><td>Thermo Fisher Scientific</td><td>BB02400600A113MNT0</td><td>24 x 60 # 1 glass cover slips</td></tr><tr><td>HCX PL APO 63x/1.3 NA Glyc</td><td>Leica</td><td>15506353</td><td>glycerol immersion objective</td></tr><tr><td>LSM 510 Meta</td><td>Zeiss</td><td>custom configuration</td><td>inverted laser scanning confocal microscope used to acquire example data 3</td></tr><tr><td>M9 Minimal Medium</td><td>Merck/Sigma-Aldrich</td><td>M6030</td><td>generic fungal growth medium</td></tr><tr><td>Micro-slide 8-well</td><td>ibidi</td><td>80826</td><td>ibiTreat #1.5 polymer coverslip</td></tr><tr><td>Miracloth</td><td>Merck/Millipore</td><td>475855-1R</td><td>polyester filtration material</td></tr><tr><td>Petri dish</td><td>Sarstedt</td><td>8,21,472</td><td>92 x 16 mm culture dish w/o cams</td></tr><tr><td>Phytagel</td><td>Merck/Sigma-Aldrich</td><td>P8169</td><td>transparent gelling agent</td></tr><tr><td>Plan Apochromat 63x/1.4 NA Oil DIC</td><td>Zeiss</td><td>440762-9904-000</td><td>oil immersion objective</td></tr><tr><td>Pontamine Fast Scarlet 4B</td><td>Merck/Sigma-Aldrich</td><td>212490</td><td>cell wall dye</td></tr><tr><td>Potato Dextrose Agar (PDA)</td><td>BD Difco</td><td>213400</td><td>fungal growth medium for &lt;em&gt;T. atroviride&lt;/em&gt;</td></tr><tr><td>Potato Dextrose Broth (PDB)</td><td>BD Difco</td><td>254920</td><td>fungal growth medium for &lt;em&gt;T. atroviride&lt;/em&gt;</td></tr><tr><td>Reaction tube</td><td>Sarstedt</td><td>72,706</td><td>1.5 mL SafeSeal tube</td></tr><tr><td>Scalpel</td><td>B.Braun</td><td>5518016</td><td>Cutfix sterile scalpel #23</td></tr><tr><td>Screw cap tube</td><td>Sarstedt</td><td>6,25,54,502</td><td>15 mL polypropylene tube</td></tr><tr><td>Solophenyl Flavine 7GFE 500</td><td>CIBA</td><td>1485385V6</td><td>cell wall dye</td></tr><tr><td>SynaptoGreen C4</td><td>Biotum</td><td>70020</td><td>membrane dye</td></tr><tr><td>SynaptoRed C2</td><td>Biotum</td><td>70021</td><td>membrane dye</td></tr><tr><td>Syringe membrane filter</td><td>Thermo Fisher Scientific</td><td>723-9945</td><td>0.45 &amp;micro;m SFCA syringe filter</td></tr><tr><td>TCS SP5 II</td><td>Leica</td><td>custom configuration</td><td>inverted laser scanning confocal microscope used to acquire example data 1</td></tr><tr><td>Vogel&#039;s Minimal Medium (VMM)</td><td>FGSC</td><td>Fungal Genetics Stock Centre</td><td>fungal growth medium for &lt;em&gt;N. crassa&lt;/em&gt;</td></tr></tbody></table>

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.

Watch this Scientific Journal Video about Application of Membrane and Cell Wall Selective Fluorescent Dyes for Live-Cell Imaging of Filamentous Fungi at JoVE.com

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

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 ...

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Nanyang Technological University

Research

JoVE Journal

Biology

21.0K Views.  University of Innsbruck. Membraan en celwand selectieve fluorescerende kleurstoffen zijn belangrijke instrumenten voor de analyse van organelle dynamiek en levende schimmelcellen. Onze protocollen bieden essentiële theoretische achtergrond en praktische richtlijnen voor de toepassing van een selectie van deze kleurstoffen als echt vitale vlekken. Het belangrijkste punt hiervoor is het gebruik van zeer lage kleurstof concentraties die niet leiden tot cellulaire artefacten in verband met verzadiging of kleurstof toxic...

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

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle synapses, we developed a larval tissue preparation that had features of live intact larvae, but also had good optical properties.  This new preparation also allowed for access of perfusates to the synapse.  We used fly larvae, immersed them in artificial hemolymph, and relaxed their normal rhythmic body contractions by chilling them. Next we dissected off the posterior segments of each animal and with a blunt insect pin pushed the mouth parts backward through the body cavity.  This everted the larval body wall, like turning a sock inside-out.  We completed the dissection with ultra-fine dissection scissors and thus exposed the visceral side of the body wall muscles. The glial structures at the NMJ expressed membrane targeted GFP under the control of glial specific promoters. The post-synaptic membrane, the SSR (Subsynaptic Reticula) in muscle expressed synaptically targeted dsRed. We needed to acutely label the motor neuron terminals, the third part of the synapse. To do this we applied primary antibodies to HRP, conjugated to a far-red emitting flurophore.  To test for dye diffusion properties into the perisynaptic space between the motor neuron terminals and the SSR, we applied a solution of large Dextran molecules conjugated to far-red emitting flurophore and collected images.

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

We described structural features of the Glia-neuromuscular synapses in a novel Inside-out tissue preparation of live fly larvae using fluorescent dyes with confocal microscopy. We labeled live neuron terminals with fluorescent primary antibodies to HRP, and also visualized the perisynaptic space with fluorescent Dextrans.

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle ...

Visualizing Yeast Organelles with Fluorescent Protein Markers

The budding yeast, Saccharomyces cerevisiae, is a classic model system in studying organelle function and dynamics. In our previous works, we have constructed fluorescent protein-based markers for major organelles and endomembrane structures, including the nucleus, endoplasmic reticulum (ER), Golgi apparatus, endosomes, vacuoles, mitochondria, peroxisomes, lipid droplets, and autophagosomes. The protocol presented here describes the procedures for using these markers in yeast, including DNA preparation for yeast transformation, selection and evaluation of transformants, fluorescent microscopic observation, and the expected outcomes. The text is geared toward researchers who are entering the field of yeast organelle study from other backgrounds. Essential steps are covered, as well as technical notes about microscope hardware considerations and several common pitfalls. It provides a starting point for people to observe yeast subcellular entities by live-cell fluorescent microscopy. These tools and methods can be used to identify protein subcellular localization and track organelles of interest in time-lapse imaging.

Here we describe the use of a set of fluorescent protein-based organelle markers in live-cell imaging of the budding yeast, Saccharomyces cerevisiae.

The budding yeast, Saccharomyces cerevisiae, is a classic model system in studying organelle function and dynamics. In our previous works, we have ...

Biochemistry

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

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

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

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

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

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 ...

Analysis of Lipid Droplet Content in Fission and Budding Yeasts using Automated Image Processing

Lipid metabolism and its regulation are of interest to both basic and applied life sciences and biotechnology. In this regard, various yeast species are used as models in lipid metabolic research or for industrial lipid production. Lipid droplets are highly dynamic storage bodies and their cellular content represents a convenient readout of the lipid metabolic state. Fluorescence microscopy is a method of choice for quantitative analysis of cellular lipid droplets, as it relies on widely available equipment and allows analysis of individual lipid droplets. Furthermore, microscopic image analysis can be automated, greatly increasing overall analysis throughput. Here, we describe an experimental and analytical workflow for automated detection and quantitative description of individual lipid droplets in three different model yeast species: the fission yeasts Schizosaccharomyces pombe and Schizosaccharomyces japonicus, and the budding yeast Saccharomyces cerevisiae. Lipid droplets are visualized with BODIPY 493/503, and cell-impermeable fluorescent dextran is added to the culture media to help identify cell boundaries. Cells are subjected to 3D epifluorescence microscopy in green and blue channels and the resulting z-stack images are processed automatically by a MATLAB pipeline. The procedure outputs rich quantitative data on cellular lipid droplet content and individual lipid droplet characteristics in a tabular format suitable for downstream analyses in major spreadsheet or statistical packages. We provide example analyses of lipid droplet content under various conditions that affect cellular lipid metabolism.

Here, we present a MATLAB implementation of automated detection and quantitative description of lipid droplets in fluorescence microscopy images of fission and budding yeast cells.

Lipid metabolism and its regulation are of interest to both basic and applied life sciences and biotechnology. In this regard, various yeast species are ...

Immunology and Infection

Live-Cell Fluorescence Microscopy to Investigate Subcellular Protein Localization and Cell Morphology Changes in Bacteria

Investigations of factors influencing cell division and cell shape in bacteria are commonly performed in conjunction with high-resolution fluorescence microscopy as observations made at a population level may not truly reflect what occurs at a single cell level. Live-cell timelapse microscopy allows investigators to monitor the changes in cell division or cell morphology which provide valuable insights regarding subcellular localization of proteins and timing of gene expression, as it happens, to potentially aid in answering important biological questions. Here, we describe our protocol to monitor phenotypic changes in Bacillus subtilis and Staphylococcus aureus using a high-resolution deconvolution microscope. The objective of this report is to provide a simple and clear protocol that can be adopted by other investigators interested in conducting fluorescence microscopy experiments to study different biological processes in bacteria as well as other organisms.

This article provides a step-by-step guide to investigate protein subcellular localization dynamics and to monitor morphological changes using high-resolution fluorescence microscopy in Bacillus subtilis and Staphylococcus aureus.

Investigations of factors influencing cell division and cell shape in bacteria are commonly performed in conjunction with high-resolution fluorescence ...

Live-cell Imaging of Fungal Cells to Investigate Modes of Entry and Subcellular Localization of Antifungal Plant Defensins

Small cysteine-rich defensins are one of the largest groups of host defense peptides present in all plants. Many plant defensins exhibit potent in vitro antifungal activity against a broad-spectrum of fungal pathogens and therefore have the potential to be used as antifungal agents in transgenic crops. In order to harness the full potential of plant defensins for diseases control, it is crucial to elucidate their mechanisms of action (MOA). With the advent of advanced microscopy techniques, live-cell imaging has become a powerful tool for understanding the dynamics of the antifungal MOA of plant defensins. Here, a confocal microscopy based live-cell imaging method is described using two fluorescently labeled plant defensins (MtDef4 and MtDef5) in combination with vital fluorescent dyes. This technique enables real-time visualization and analysis of the dynamic events of MtDef4 and MtDef5 internalization into fungal cells. Importantly, this assay generates a wealth of information including internalization kinetics, mode of entry and subcellular localization of these peptides. Along with other cell biological tools, these methods have provided critical insights into the dynamics and complexity of the MOA of these peptides. These tools can also be used to compare the MOA of these peptides against different fungi.

Plant defensins play an important role in plant defense against pathogens. For effective use of these antifungal peptides as antifungal agents, understanding their modes of action (MOA) is critical. Here, a live-cell imaging method is described to study critical aspects of the MOA of these peptides.

Small cysteine-rich defensins are one of the largest groups of host defense peptides present in all plants. Many plant defensins exhibit potent in vitro ...

Live Cell Fluorescence Microscopy to Observe Essential Processes During Microbial Cell Growth

Core cellular processes such as DNA replication and segregation, protein synthesis, cell wall biosynthesis, and cell division rely on the function of proteins which are essential for bacterial survival. A series of target-specific dyes can be used as probes to better understand these processes. Staining with lipophilic dyes enables the observation of membrane structure, visualization of lipid microdomains, and detection of membrane blebs. Use of fluorescent-d-amino acids (FDAAs) to probe the sites of peptidoglycan biosynthesis can indicate potential defects in cell wall biogenesis or cell growth patterning. Finally, nucleic acid stains can indicate possible defects in DNA replication or chromosome segregation. Cyanine DNA stains label living cells and are suitable for time-lapse microscopy enabling real-time observations of nucleoid morphology during cell growth. Protocols for cell labeling can be applied to protein depletion mutants to identify defects in membrane structure, cell wall biogenesis, or chromosome segregation. Furthermore, time-lapse microscopy can be used to monitor morphological changes as an essential protein is removed and can provide additional insights into protein function. For example, the depletion of essential cell division proteins results in filamentation or branching, whereas the depletion of cell growth proteins may cause cells to become shorter or rounder. Here, protocols for cell growth, target-specific labeling, and time-lapse microscopy are provided for the bacterial plant pathogen Agrobacterium tumefaciens. Together, target-specific dyes and time-lapse microscopy enable characterization of essential processes in A. tumefaciens. Finally, the protocols provided can be readily modified to probe essential processes in other bacteria.

Understanding the function of essential processes in bacteria is challenging. Fluorescence microscopy with target-specific dyes can provide key insights into microbial cell growth and cell cycle progression. Here, Agrobacterium tumefaciens is used as a model bacterium to highlight methods for live cell imaging for characterization of essential processes.

Core cellular processes such as DNA replication and segregation, protein synthesis, cell wall biosynthesis, and cell division rely on the function of ...

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 ...

Examination of Mitotic and Meiotic Fission Yeast Nuclear Dynamics by Fluorescence Live-cell Microscopy

Live-cell imaging is a microscopy technique used to examine cell and protein dynamics in living cells. This imaging method is not toxic, generally does not interfere with cell physiology, and requires minimal experimental handling. The low levels of technical interference enable researchers to study cells across multiple cycles of mitosis and to observe meiosis from beginning to end. Using fluorescent tags such as Green Fluorescent Protein (GFP) and Red Fluorescent Protein (RFP), researchers can analyze different factors whose functions are important for processes like transcription, DNA replication, cohesion, and segregation. Coupled with data analysis using Fiji (a free, optimized ImageJ version), live-cell imaging offers various ways of assessing protein movement, localization, stability, and timing, as well as nuclear dynamics and chromosome segregation. However, as is the case with other microscopy methods, live-cell imaging is limited by the intrinsic properties of light, which put a limit to the resolution power at high magnifications, and is also sensitive to photobleaching or phototoxicity at high wavelength frequencies. However, with some care, investigators can bypass these physical limitations by carefully choosing the right conditions, strains, and fluorescent markers to allow for the appropriate visualization of mitotic and meiotic events.

Here, we present live-cell imaging which is a non-toxic microscopy method that allows researchers to study protein behavior and nuclear dynamics in living fission yeast cells during mitosis and meiosis.

Live-cell imaging is a microscopy technique used to examine cell and protein dynamics in living cells. This imaging method is not toxic, generally does ...

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

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