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 T. atroviride</td>
		</tr>
		<tr>
			<td>Potato Dextrose Broth (PDB)</td>
			<td>BD Difco</td>
			<td>254920</td>
			<td>fungal growth medium for T. atroviride</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 &#181;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&#39;s Minimal Medium (VMM)</td>
			<td>FGSC</td>
			<td>Fungal Genetics Stock Centre</td>
			<td>fungal growth medium for N. crassa</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 ...

application-membrane-cell-wall-selective-fluorescent-dyes-for-live

Nanyang Technological University

Research

JoVE Journal

Biology

20.7K Views.  University of Innsbruck. 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.

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

Labeling Stem Cells with Fluorescent Dyes for non-invasive Detection with Optical Imaging

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo labeling of stem cells with a fluorescent dye, subsequent intravenous injection of the labeled cells and visualization of their accumulation in specific target organs or pathologies. The presented technique demonstrates how we label human mesenchymal stem cells (hMSC) by simple incubation with the lipophilic fluorescent dye DiD (C67H103CIN2O3S) and how we label human embryonic stem cells (hESC) with the FDA approved fluorescent dye Indocyanine Green (ICG). The uptake mechanism is via adherence and diffusion of the lypophilic dye across the phospholipid cell membrane bilayer. The labeling efficiency is usually improved if the cells are incubated with the dye in serum-free media as opposed to incubation in serum-containing media. Furthermore, the addition of the transfection agent Protamine Sulfate significantly improves contrast agent uptake.

This video shows techniques for labeling of human embryonic stem cells and mesenchymal stem cells with fluorescent dyes. This technique can be used for an in vivo tracking of transplanted stem cells with optical imaging and for histopathological correlations with fluorescence microscopy.

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo ...

Selective Labelling of Cell-surface Proteins using CyDye DIGE Fluor Minimal Dyes

Surface  proteins are central to the cell's ability to react to its environment and to interact with neighboring cells. They are known to be inducers of almost all intracellular signaling. Moreover, they play an important role in environmental adaptation and drug treatment, and are often involved in disease pathogenesis and pathology (1). Protein-protein interactions are intrinsic to signaling pathways, and to gain more insight in these complex biological processes, sensitive and reliable methods are needed for studying cell surface proteins. Two-dimensional (2-D) electrophoresis is used extensively for detection of biomarkers and other targets in complex protein samples to study differential changes. Cell surface proteins, partly due to their low abundance (1 2% of cellular proteins), are difficult to detect in a 2-D gel without fractionation or some other type of enrichment. They are also often poorly represented in 2-D gels due to their hydrophobic nature and high molecular weight (2). In this study, we present a new protocol for intact cells using CyDye  DIGE Fluor minimal dyes for specific labeling and detection of this important group of proteins. The results showed specific labeling of a large number of cell surface proteins with minimal labeling of intracellular proteins. This protocol is rapid, simple to use, and all three CyDye DIGE Fluor minimal dyes (Cy 2, Cy 3 and Cy 5) can be used to label cell-surface proteins. These features allow for multiplexing using the 2-D Fluorescence Difference Gel Electrophoresis (2-D DIGE) with Ettan  DIGE technology and analysis of protein expression changes using DeCyder  2-D Differential Analysis Software. The level of cell-surface proteins was followed during serum starvation of CHO cells for various lengths of time (see Table 1). Small changes in abundance were detected with high accuracy, and results are supported by defined statistical methods.

A simple and specific method was demonstrated for fluorescent labeling and enhanced detection of cell surface proteins without a fractionation step. Differential abundance in cell surface proteins was analyzed using two-dimensional (2-D) electrophoresis and Ettan&#x2122; DIGE technology.

Surface  proteins are central to the cell's ability to react to its environment and to interact with neighboring cells. They are known to be inducers of ...

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy (FSM)

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique advantage of FSM is its ability to capture the movement and turnover kinetics (assembly/disassembly) of the F-actin network within living cells. This technique is particularly useful in deriving quantitative measurements of F-actin dynamics when paired with computer vision software (qFSM).  We describe  the selection, microinjection and visualization of fluorescent actin probes in living cells. Importantly, similar procedures are applicable to visualizing other macomolecular assemblies. FSM has been demonstrated for microtubules, intermediate filaments, and adhesion complexes.  

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy FSM

Selection, microinjection, and imaging of fluorescently-labeled F-actin via fluorescent speckle microscopy (FSM).

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique ...

Fluorescent Labeling of COS-7 Expressing SNAP-tag Fusion Proteins for Live Cell Imaging

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. These systems offer a broad selection of fluorescent substrates optimized for a range of imaging instrumentation. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again. There are two steps to using this system: cloning and expression of the protein of interest as a SNAP-tag fusion, and labeling of the fusion with the SNAP-tag substrate of choice. The SNAP-tag is a small protein based on human O6-alkylguanine-DNA-alkyltransferase (hAGT), a DNA repair protein. SNAP-tag labels are dyes conjugated to guanine or chloropyrimidine leaving groups via a benzyl linker. In the labeling reaction, the substituted benzyl group of the substrate is covalently attached to the SNAP-tag. CLIP-tag is a modified version of SNAP-tag, engineered to react with benzylcytosine rather than benzylguanine derivatives. When used in conjunction with SNAP-tag, CLIP-tag enables the orthogonal and complementary labeling of two proteins simultaneously in the same cells.

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again.

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest ...

Two- and Three-Dimensional Live Cell Imaging of DNA Damage Response Proteins

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate mutations and chromosomal aberrations1. To prevent this trauma from catalyzing genomic instability, it is crucial for cells to detect DSBs, activate the DNA damage response (DDR), and repair the DNA. When stimulated, the DDR works to preserve genomic integrity by triggering cell cycle arrest to allow for repair to take place or force the cell to undergo apoptosis. The predominant mechanisms of DSB repair occur through nonhomologous end-joining (NHEJ) and homologous recombination repair (HRR) (reviewed in2). There are many proteins whose activities must be precisely orchestrated for the DDR to function properly. Herein, we describe a method for 2- and 3-dimensional (D) visualization of one of these proteins, 53BP1.
The p53-binding protein 1 (53BP1) localizes to areas of DSBs by binding to modified histones3,4, forming foci within 5-15 minutes5. The histone modifications and recruitment of 53BP1 and other DDR proteins to DSB sites are believed to facilitate the structural rearrangement of chromatin around areas of damage and contribute to DNA repair6. Beyond direct participation in repair, additional roles have been described for 53BP1 in the DDR, such as regulating an intra-S checkpoint, a G2/M checkpoint, and activating downstream DDR proteins7-9. Recently, it was discovered that 53BP1 does not form foci in response to DNA damage induced during mitosis, instead waiting for cells to enter G1 before localizing to the vicinity of DSBs6. DDR proteins such as 53BP1 have been found to associate with mitotic structures (such as kinetochores) during the progression through mitosis10.
In this protocol we describe the use of 2- and 3-D live cell imaging to visualize the formation of 53BP1 foci in response to the DNA damaging agent camptothecin (CPT), as well as 53BP1's behavior during mitosis. Camptothecin is a topoisomerase I inhibitor that primarily causes DSBs during DNA replication. To accomplish this, we used a previously described 53BP1-mCherry fluorescent fusion protein construct consisting of a 53BP1 protein domain able to bind DSBs11. In addition, we used a histone H2B-GFP fluorescent fusion protein construct able to monitor chromatin dynamics throughout the cell cycle but in particular during mitosis12. Live cell imaging in multiple dimensions is an excellent tool to deepen our understanding of the function of DDR proteins in eukaryotic cells.

This protocol describes a method for visualizing a DNA double-strand break signaling protein activated in response to DNA damage as well as its localization during mitosis.

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate ...

Live Cell Imaging of Early Autophagy Events: Omegasomes and Beyond

Autophagy is a cellular response triggered by the lack of nutrients, especially the absence of amino acids. Autophagy is defined by the formation of double membrane structures, called autophagosomes, that sequester cytoplasm, long-lived proteins and protein aggregates, defective organelles, and even viruses or bacteria. Autophagosomes eventually fuse with lysosomes leading to bulk degradation of their content, with the produced nutrients being recycled back to the cytoplasm. Therefore, autophagy is crucial for cell homeostasis, and dysregulation of autophagy can lead to disease, most notably neurodegeneration, ageing and cancer.	
	Autophagosome formation is a very elaborate process, for which cells have allocated a specific group of proteins, called the core autophagy machinery. The core autophagy machinery is functionally complemented by additional proteins involved in diverse cellular processes, e.g. in membrane trafficking, in mitochondrial and lysosomal biology. Coordination of these proteins for the formation and degradation of autophagosomes constitutes the highly dynamic and sophisticated response of autophagy. Live cell imaging allows one to follow the molecular contribution of each autophagy-related protein down to the level of a single autophagosome formation event and in real time, therefore this technique offers a high temporal and spatial resolution.	
	Here we use a cell line stably expressing GFP-DFCP1, to establish a spatial and temporal context for our analysis. DFCP1 marks omegasomes, which are precursor structures leading to autophagosomes formation. A protein of interest (POI) can be marked with either a red or cyan fluorescent tag. Different organelles, like the ER, mitochondria and lysosomes, are all involved in different steps of autophagosome formation, and can be marked using a specific tracker dye. Time-lapse microscopy of autophagy in this experimental set up, allows information to be extracted about the fourth dimension, i.e. time. Hence we can follow the contribution of the POI to autophagy in space and time.

Time-lapse microscopy of fluorescently labeled autophagy markers allows monitoring of the dynamic autophagy response with high temporal resolution. Using specific autophagy and organelle markers in a combination of 3 different colors, we can follow the contribution of a protein to autophagosome formation in a robust spatial and temporal context.

Autophagy is a  cellular response triggered by the lack of nutrients, especially the absence of  amino acids. Autophagy is defined by the formation of ...

Intravital Microscopy for Imaging Subcellular Structures in Live Mice Expressing Fluorescent Proteins

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, we use the salivary glands of live mice since they provide several advantages. First, they can be easily exposed to enable access to the optics, and stabilized to facilitate the reduction of the motion artifacts due to heartbeat and respiration. This significantly facilitates imaging and tracking small subcellular structures. Second, most of the cell populations of the salivary glands are accessible from the surface of the organ. This permits the use of confocal microscopy that has a higher spatial resolution than other techniques that have been used for in vivo imaging, such as two-photon microscopy. Finally, salivary glands can be easily manipulated pharmacologically and genetically, thus providing a robust system to investigate biological processes at a molecular level.
In this study we focus on a protocol designed to follow the kinetics of the exocytosis of secretory granules in acinar cells and the dynamics of the apical plasma membrane where the secretory granules fuse upon stimulation of the beta-adrenergic receptors. Specifically, we used a transgenic mouse that co-expresses cytosolic GFP and a membrane-targeted peptide fused with the fluorescent protein tandem-Tomato. However, the procedures that we used to stabilize and image the salivary glands can be extended to other mouse models and coupled to other approaches to label in vivo cellular components, enabling the visualization of various subcellular structures, such as endosomes, lysosomes, mitochondria, and the actin cytoskeleton.

Intravital microscopy is a powerful tool that enables imaging various biological processes in live animals. In this article, we present a detailed method for imaging the dynamics of subcellular structures, such as the secretory granules, in the salivary glands of live mice.

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, ...

G Protein-selective GPCR Conformations Measured Using FRET Sensors in a Live Cell Suspension Fluorometer Assay

F&#1255;rster resonance energy transfer (FRET)-based studies have become increasingly common in the investigation of GPCR signaling. Our research group developed an intra-molecular FRET sensor to detect the interaction between G&#945; subunits and GPCRs in live cells following agonist stimulation. Here, we detail the protocol for detecting changes in FRET between the &#946;2-adrenergic receptor and the G&#945;s C-terminus peptide upon treatment with 100 &#181;M isoproterenol hydrochloride as previously characterized1. Our FRET sensor is a single polypeptide consisting serially of a full-length GPCR, a FRET acceptor fluorophore (mCitrine), an ER/K SPASM (systematic protein affinity strength modulation) linker, a FRET donor fluorophore (mCerulean), and a G&#945; C-terminal peptide. This protocol will detail cell preparation, transfection conditions, equipment setup, assay execution, and data analysis. This experimental design detects small changes in FRET indicative of protein-protein interactions, and can also be used to compare the strength of interaction across ligands and GPCR-G protein pairings. To enhance the signal-to-noise in our measurements, this protocol requires heightened precision in all steps, and is presented here to enable reproducible execution.

Simple methods to detect the selective activation of G proteins by G protein-coupled receptors remain an outstanding challenge in cell signaling. Here, F&#1255;rster resonance energy transfer (FRET) biosensors have been developed by pairwise tethering a GPCR to G protein peptides to probe conformational changes at controlled concentrations in live cells.

F&#1255;rster resonance energy transfer (FRET)-based studies have become increasingly common in the investigation of GPCR signaling. Our research group ...

Studying the Protein Quality Control System of D. discoideum Using Temperature-controlled Live Cell Imaging

The complex lifestyle of the social amoebae Dictyostelium discoideum makes it a valuable model for the study of various biological processes. Recently, we showed that D. discoideum is remarkably resilient to protein aggregation and can be used to gain insights into the cellular protein quality control system. However, the use of D. discoideum as a model system poses several challenges to microscopy-based experimental approaches, such as the high motility of the cells and their susceptibility to photo-toxicity. The latter proves to be especially challenging when studying protein homeostasis, as the phototoxic effects can induce a cellular stress response and thus alter to behavior of the protein quality control system. 
Temperature increase is a commonly used way to induce cellular stress. Here, we describe a temperature-controllable imaging protocol, which allows observing temperature-induced perturbations in D. discoideum. Moreover, when applied at normal growth temperature, this imaging protocol can also noticeably reduce photo-toxicity, thus allowing imaging with higher intensities. This can be particularly useful when imaging proteins with very low expression levels. Moreover, the high mobility of the cells often requires the acquisition of multiple fields of view to follow individual cells, and the number of fields needs to be balanced against the desired time interval and exposure time.

Studying the Protein Quality Control System of D. discoideum Using Temperature-controlled Live Cell Imaging

The social amoebae Dictyostelium discoideum has recently been established as a system to study protein misfolding and proteostasis. Here, we describe a new imaging-based methodology to study temperature-induced protein aggregation and the cellular stress response in D. discoideum.

The complex lifestyle of the social amoebae Dictyostelium discoideum makes it a valuable model for the study of various biological processes. Recently, we ...

Live-cell Imaging of Platelet Degranulation and Secretion Under Flow

Blood platelets are essential players in hemostasis, the formation of thrombi to seal vascular breaches. They are also involved in thrombosis, the formation of thrombi that occlude the vasculature and injure organs, with life-threatening consequences. This motivates scientific research on platelet function and the development of methods to track cell-biological processes as they occur under flow conditions.
A variety of flow models are available for the study of platelet adhesion and aggregation, two key phenomena in platelet biology. This work describes a method to study real-time platelet degranulation under flow during activation. The method makes use of a flow chamber coupled to a syringe-pump setup that is placed under a wide-field, inverted, LED-based fluorescence microscope. The setup described here allows for the simultaneous excitation of multiple fluorophores that are delivered by fluorescently labeled antibodies or fluorescent dyes. After live-cell imaging experiments, the cover glasses can be further processed and analyzed using static microscopy (i.e., confocal microscopy or scanning electron microscopy).

This work describes a fluorescence microscopy-based method for the study of platelet adhesion, spreading, and secretion under flow. This versatile platform enables the investigation of platelet function for mechanistic research on thrombosis and hemostasis.