We thank Dr. R. Howard Berg, Director of the Integrated Microscopy Facility at the Donald Danforth Plant Science Center, for his guidance and help with confocal microscopy. The authors have no conflict of interest to declare.

1. Labeling of Defensins with Fluorophores
<ol>
	<li>Select a fluorophore that has minimum effect on the antimicrobial properties as well as the uptake and localization of defensin inside the living cell. 
	NOTE: Selecting optimal fluorophore depends on specific experimental objectives. The spectral and chemical properties, photostability, size and charge of the fluorophore should also be considered.</li>
	<li>Label defensin with the selected fluorophore using an appropriate peptide labeling kit available from commercial vendors. Alternatively, chemically synthesize fluorophore-labeled defensins. In this study, label MtDef5 defensin with DyLight550 using a commercial labeling kit according to manufacturer&#39;s protocol and tetramethyl rhodamine (TAMRA)-labeled MtDef4 defensin was chemically synthesized commercially. 
	NOTE: It is recommended that chemically synthesized fluorophore tagged peptides (especially for short peptides of 50 amino acids or less) be used since the fluorophore can be attached to the N- or C-terminal residue of the peptide to ensure minimum effect on its antimicrobial activity. Synthesis of long peptides with more than three disulfide bonds is challenging. In this case, it is recommended that peptide be labeled with an optimal fluorophore using a suitable commercially available peptide labeling kit.</li>
	<li>Test in vitro antimicrobial activity of the labeled defensin and determine the minimal inhibitory concentration (MIC) against the fungi to be investigated. 
	NOTE: In this study, the MIC of DyLight550-MtDef5 and TMR-MtDef4 was determined against Neurospora crassa and Fusarium graminearum.</li>
</ol>
2. Fungal Cultures and Growth Medium
<ol>
	<li>Transfer conidia from a N. crassa stock culture to slant tube containing Vogel&#39;s agar medium21 and incubate at room temperature under light for 5 days.</li>
	<li>Culture F. graminearum PH-1 strain on plates containing complete medium (CM)22 at 28 &#186;C for 5 days. For production of conidia, inoculate 4 (10 mm diameter) plugs of 5 day old culture of F. graminearum PH-1 into 50 mL of carboxymethyl cellulose (CMC) medium (15 g carboxymethyl cellulose, 1 g Yeast extract, 0.5 g MgSO4.7 H2O, 1 g NH4NO3, and 1 g KH2PO4) and culture for 4-7 days at 28 &#186;C in a rotary shaker at 180 rpm.</li>
</ol>
3. Preparation of Conidial Suspension
<ol>
	<li>F. graminearum conidial suspension
	<ol>
		<li>Vortex the F. graminearum liquid culture and collect conidia by filtering 1.5 mL fungal culture through two layers of filtration material (such as Miracloth)&#160;into a 2 mL microcentrifuge tube. Centrifuge the conidial suspension at 13,226 x g speed in a microcentrifuge for 2 min.</li>
		<li>Discard the supernatant, and add 1 mL of sterile water to wash the pellet.</li>
		<li>Centrifuge the conidial suspension at 13,226 x g speed in a microcentrifuge for 2 min. Discard the supernatant and resuspend the pellet in 1 mL of 2X SFM (Synthetic Fungal Media)23.</li>
		<li>Count the conidia using a hemocytometer under a light microscope.</li>
		<li>Adjust the conidial suspension to 105 conidia/mL with 2X SFM.</li>
	</ol>
	</li>
	<li>N. crassa conidial suspension
	<ol>
		<li>Transfer a small amount of growing culture (5 day old) of N. crassa using an inoculation loop to a microcentrifuge tube containing 2 mL of Vogel&#39;s liquid medium.</li>
		<li>Vortex the conidial suspension and filter through filtration material into a new microcentrifuge tube.</li>
		<li>Centrifuge the conidial suspension at maximum speed in a microcentrifuge for 2 min. Discard the supernatant and resuspend the conidial pellet in 1 mL of Vogel&#39;s liquid medium.</li>
		<li>Count the conidia using a hemocytometer under a light microscope.</li>
		<li>Adjust the conidial suspension to 105 conidia/mL with Vogel&#39;s liquid medium.</li>
	</ol>
	</li>
</ol>
4. Sample Preparation and Confocal Microscopy
<ol>
	<li>Subcellular localization of MtDef4 defensin into fungal cells
	<ol>
		<li>Pipette 50 &#181;L of each conidial suspension (105 conidia/mL) into the 10 mm microwell of 35 mm glass bottom microwell dishes. For the observation of conidia, directly proceed to the next step, or for germlings preparation, pipette 50 &#181;L of N. crassa and F. graminearum conidia into the 35 mm culture dishes and let germinate for 3-6 h at room temperature.</li>
		<li>Add 50 &#181;L of fluorescently labeled defensin at the MIC to each microwell dish and incubate for 2.5 h. The MIC of the fluorophore-labeled defensins used here (step 1.3) is 3 &#181;M for both N. crassa and F. graminearum and the MIC of TMR-MtDef4 for N. crassa and F. graminearum was 1 &#181;M and 12 &#181;M, respectively.</li>
		<li>Add 2 &#181;L of membrane-selective dye FM4-64 (final concentration: 5 &#181;M) in culture dish and incubate for 30 min and mount immediately on the confocal microscope for imaging.</li>
		<li>Select the White Light laser (WLL). Use the two laser sources 488 nm and 550 nm to excite TMR-MtDef4 and FM4-64 dye, respectively. Detect fluorescence of TMR-MtDef4 at 580-700 nm and detect FM4-64 dye at 690-800 nm. 
		NOTE: Perform confocal microscopy in the dark room.</li>
	</ol>
	</li>
	<li>Time lapse imaging of internalization of MtDef5 defensin
	<ol>
		<li>Pipette 50 &#181;L of N. crassa and F. graminearum conidial suspension (105 conidia/mL) into 10 mm microwell of 35 mm glass bottom microwell dishes. The 10 mm microwell is uncoated and has a No. 1.5 cover glass at the bottom. For the observation of conidia, directly proceed to the next step. For observation of germlings, incubate conidia for 3-6 h at room temperature before proceeding with the microscopy.</li>
		<li>Turn on WLL. Select the laser line at 550 nm to excite the fluorophore-labeled defensins used here (step 1.3) and FM4-64 with 1.00% intensity and activate the corresponding detectors. The fluorophore-labeled defensins used here (step 1.3) was detected at 560 - 600 nm and the FM4-64dye was detected at 690 - 800 nm. 
		NOTE: Use low laser intensity for live cell imaging as high laser intensity can cause damage to the live fungal cells.</li>
		<li>Mount the microwell dish on the microscope. Find the cells using 10 x objectives initially, then switch to 100x/1.44 OIL objective for higher magnification. 
		NOTE: Set the scan mode to xyzt (combination of Z-stack and time-lapse modes), Z position, Zoom, frequency of image capture, etc. before proceeding to the next step.</li>
		<li>Add 50 &#181;L of the fluorophore-labeled defensins used here (step 1.3) of at the MIC of 3 &#181;M and 2 &#181;L of membrane-selective dye FM4-64 (final concentration 5 &#181;M) to each microwell dish, and start time-lapse imaging. Place small pieces of wet filter papers in the microwell dishes to prevent evaporation. The frequency of image capture was 3 min 30 s and the total period was 2 h 30 min. 
		NOTE: Adding defensin and fluorophore after mounting microwell dish in the microscope can defocus the region of interest. Therefore, after adding defensin and fluorophore to the microwell, region of interest needs to be refocused which can cause delay in image acquisition.</li>
		<li>Use a confocal microscope for all confocal imaging and carry out the microscopy at room temperature in a dark room.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

In this study, a reliable live-cell imaging methodology with the use of fluorescently labeled antifungal defensins was described to study the kinetics of the internalization of these peptides into fungal cells and to determine their subcellular targets. This method is a powerful tool to visualize the dynamics of the interaction between defensins and fungal cells temporally and spatially.
Various methods have been used to study the internalization and intracellular localization of plant defensi...

Plants have evolved a sophisticated innate immune system for defense against the microbial plant pathogens1. They express numerous gene-encoded host defense peptides with putative antimicrobial activity2. Indeed, many of these peptides display antimicrobial activity in vitro3. Defensins comprise one of the largest groups of host defense peptides in the plant kingdom4. These cysteine-rich, cationic peptides exhibit potent growth inhibitory activity against fungal and oomycete pathogens at micromolar concentrations and represent one of the first lines of defense against these pathogens5,6. Because of their potent antifungal activity, defensins can be exploited in agribiotechnological applications to generate disease resistant crops. Constitutive overexpression of several plant defensins has been shown to enhance disease resistance in the greenhouse and field tests of transgenic crops6. It is important to unravel the mechanisms of action (MOA) of these antifungal peptides in order to fully harness their potential as effective tools for crop protection. However, the MOA of these plant defensins are poorly understood. Current evidence suggests that they exhibit different MOA5,6,7,8. Some defensins act extracellularly on fungi and target specific cell wall/plasma membrane resident sphingolipids, disrupt membrane integrity and activate cellular toxicity pathways9,10,11. Recently, however, antifungal defensins that translocate into fungal cells have been discovered12,13,14. Some of these defensins bind to membrane-resident bioactive phospholipids, form oligomeric complexes and permeabilize plasma membranes15,16,17. Thus, some aspects of the MOA of plant defensins have been elucidated. However, the MOA of plant defensins likely involve a complex set of events which have not yet been identified and integrated into a comprehensive model. In particular, there remains a major gap in our understanding of the cellular targets of these peptides.
With recent advances in microscopy technologies and the development of new fluorescent probes, live-cell imaging techniques are now frequently used to study the MOA of antimicrobial peptides (AMPs). These techniques complement widely used methods such as immunolocalization, electron microscopy, atomic force microscopy or X-ray tomography18, which have been employed mostly to analyze the effects of antifungal peptides on the morphology and growth of fungal cells including the study of cell wall integrity, alterations in cell growth/branching patterns, as well as plasma membrane permeabilization and killing. Nevertheless, these studies have been limited to imaging cells at a certain time point after treatment with the peptides instead of performing time-lapse imaging on the same cells to monitor their dynamic changes in response to defensin challenge. In recent years, use of fluorescently labeled peptides in conjunction with live cell imaging using confocal microscopy has enabled real-time visualization of the dynamics of AMP&#8211;microbe interactions. Both naturally purified and chemically synthesized antifungal peptides can be tagged with fluorescent labels (e.g., DyLight, rhodamine, BODIPY, or Alexa Fluor based dyes) and directly observed during their interaction with cells by time-lapse live-cell imaging. The use of these labeled peptides has significantly increased our understanding of the different aspects of their MOA including mode of entry, subcellular localization, intracellular trafficking, and sites of antifungal action within living fungal cells18.
Recently, several studies have shown that various antifungal peptides including plant defensins are internalized by living fungal cells12,14,19,20. The MOA of these defensins likely involve interaction with intracellular targets. We have recently reported the antifungal action of a plant defensin MtDef4 in two ascomycete fungi, Neurospora crassa and Fusarium graminearum. MtDef4 was shown to use different pathways for fungal cell entry and subcellular localization in these fungi14. This study used chemically synthesized tetramethyl rhodamine (TAMRA)-labeled MtDef4 in combination with vital fluorescent dyes (the membrane selective dye, FM4-64; membrane-permeant dye, SYTOX Green; the cell death reporter dye, propidium iodide) and metabolic inhibitors. These analyses demonstrated the kinetics of the internalization of MtDef4, its mechanisms of intracellular transportation and its subcellular targets14.
Here, a protocol for live-cell imaging using confocal microscopy is presented. The protocol utilizes fluorescently labeled peptides in combination with vital fluorescent dyes to study plant defensin-fungal interactions, in particular, the pathways of translocation and the intracellular targets of defensins in fungal cells.

<table><tbody><tr><td>FM4-64 Dye</td><td>Life Technologies</td><td>T13320</td><td></td></tr><tr><td>DyLight 550 Antibody Labeling Kit</td><td>Thermo Scientific</td><td>84530</td><td></td></tr><tr><td>Glass Bottom Microwell Dishes</td><td>Mat TeK</td><td>P35G-1.5-10-C</td><td></td></tr><tr><td>Mira cloth</td><td>EMD Millipore Corp</td><td>475855-1R</td><td></td></tr><tr><td>SP8-X confocal microscope</td><td>Leica</td><td></td><td></td></tr><tr><td>ImageJ software</td><td>FiJi</td><td></td><td>For Image analysis</td></tr><tr><td>Imaris software</td><td>Bitplane</td><td></td><td>For Image analysis</td></tr></tbody></table>

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.

Live cell imaging was carried out to track and compare the internalization and subcellular localization of two defensins, MtDef4 and MtDef5, from Medicago truncatula; in fungal cells. TMR-MtDef4 was chemically synthesized while MtDef5 was labeled with Dylight550 (Dylight550-MtDef5). Conidia were incubated with either defensin in combination with the membrane selective dye FM4-64. Figure 1 shows that TMR-MtDef4 has different trafficking pathways in <e...

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

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 ...

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Research

JoVE Journal

Biology

6.6K Views.  Donald Danforth Plant Science Center. 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.

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

Live-cell Video Microscopy of Fungal Pathogen Phagocytosis

Phagocytic clearance of fungal pathogens, and microorganisms more generally, may be considered to consist of four distinct stages: (i) migration of phagocytes to the site where pathogens are located; (ii) recognition of pathogen-associated molecular patterns (PAMPs) through pattern recognition receptors (PRRs); (iii) engulfment of microorganisms bound to the phagocyte cell membrane, and (iv) processing of engulfed cells within maturing phagosomes and digestion of the ingested particle. Studies that assess phagocytosis in its entirety are informative1, 2, 3, 4, 5 but are limited in that they do not normally break the process down into migration, engulfment and phagosome maturation, which may be affected differentially. Furthermore, such studies assess uptake as a single event, rather than as a continuous dynamic process. We have recently developed advanced live-cell imaging technologies, and have combined these with genetic functional analysis of both pathogen and host cells to create a cross-disciplinary platform for the analysis of innate immune cell function and fungal pathogenesis. These studies have revealed novel aspects of phagocytosis that could only be observed using systematic temporal analysis of the molecular and cellular interactions between human phagocytes and fungal pathogens and infectious microorganisms more generally. For example, we have begun to define the following: (a) the components of the cell surface required for each stage of the process of recognition, engulfment and killing of fungal cells1, 6, 7, 8; (b) how surface geometry influences the efficiency of macrophage uptake and killing of yeast and hyphal cells7; and (c) how engulfment leads to alteration of the cell cycle and behavior of macrophages 9, 10.
In contrast to single time point snapshots, live-cell video microscopy enables a wide variety of host cells and pathogens to be studied as continuous sequences over lengthy time periods, providing spatial and temporal information on a broad range of dynamic processes, including cell migration, replication and vesicular trafficking. Here we describe in detail how to prepare host and fungal cells, and to conduct the video microscopy experiments. These methods can provide a user-guide for future studies with other phagocytes and microorganisms.

We describe methods for live-cell video microscopy of Candida albicans phagocytosis by macrophages. These methods enable stage-specific analysis of macrophage migration, recognition, engulfment and phagosome maturation and reveal novel aspects of phagocytosis.

Phagocytic clearance of fungal pathogens, and microorganisms more generally, may be considered to consist of four distinct stages: (i) migration of ...

Immunology and Infection

Live-cell Imaging and Quantitative Analysis of Embryonic Epithelial Cells in Xenopus laevis

Embryonic epithelial cells serve as an ideal model to study morphogenesis where multi-cellular tissues undergo changes in their geometry, such as changes in cell surface area and cell height, and where cells undergo mitosis and migrate. Furthermore, epithelial cells can also regulate morphogenetic movements in adjacent tissues1. A traditional method to study epithelial cells and tissues involve chemical fixation and histological methods to determine cell morphology or localization of particular proteins of interest. These approaches continue to be useful and provide "snapshots" of cell shapes and tissue architecture, however, much remains to be understood about how cells acquire specific shapes, how various proteins move or localize to specific positions, and what paths cells follow toward their final differentiated fate. High resolution live imaging complements traditional methods and also allows more direct investigation into the dynamic cellular processes involved in the formation, maintenance, and morphogenesis of multicellular epithelial sheets. Here we demonstrate experimental methods from the isolation of animal cap tissues from Xenopus laevis embryos to confocal imaging of epithelial cells and simple measurement approaches that together can augment molecular and cellular studies of epithelial morphogenesis.

Live-cell Imaging and Quantitative Analysis of Embryonic Epithelial Cells in Xenopus laevis

Xenopus embryonic epithelia are an ideal model system to study cell behaviors such as polarity development and shape change during epithelial morphogenesis. Traditional histology of fixed samples is increasingly being complemented by live-cell confocal imaging. Here we demonstrate methods to isolate frog tissues and visualize live epithelial cells and their cytoskeleton using live-cell confocal microscopy.

Embryonic epithelial cells serve as an ideal model to study morphogenesis where multi-cellular tissues undergo changes in their geometry, such as changes ...

Live Imaging of Antifungal Activity by Human Primary Neutrophils and Monocytes in Response to A. fumigatus

Aspergillus fumigatus is an opportunistic fungal pathogen causing invasive infections in immunocompromised hosts with a high case-fatality rate. Research investigating immunological responses against A. fumigatus has been limited by the lack of consistent and reliable assays for measuring the antifungal activity of specific immune cells in vitro. A new method is described to assess the antifungal activity of primary monocytes and neutrophils from human donors against A. fumigatus using FLuorescent Aspergillus REporter (FLARE) conidia. These conidia contain a genetically encoded dsRed reporter, which is constitutively expressed by live FLARE conidia, and are externally labeled with Alexa Fluor 633, which is resistant to degradation within the phagolysosome, thus allowing a distinction between live and dead A. fumigatus conidia. Video microscopy and flow cytometry are subsequently used to visualize the interaction between conidia and innate immune cells, assessing fungicidal activity whilst also providing a wealth of information on phagocyte migration, phagocytosis and the inhibition of fungal growth. This novel technique has already provided exciting new insights into the host-pathogen interaction of primary immune cells against A. fumigatus. It is important to note the laboratory setup required to perform this assay, including the necessary microscopy and flow cytometry facilities, and the capacity to work with human donor blood and genetically manipulated fungi. However, this assay is capable of generating large amounts of data and can reveal detailed insights into the antifungal response. This protocol has successfully been used to study the host-pathogen interaction of primary immune cells against A. fumigatus.
It is important to note the laboratory setup required to perform this assay, including the necessary microscopy and flow cytometry facilities, and the capacity to work with human donor blood and genetically manipulated fungi. However, this assay is capable of generating large amounts of data and can reveal detailed insights into the antifungal response. This protocol has successfully been used to study the host-pathogen interaction of primary immune cells against A. fumigatus.

Live Imaging of Antifungal Activity by Human Primary Neutrophils and Monocytes in Response to A. fumigatus

Here, we describe a protocol to assess antifungal activity of primary human immune cells in real-time using fluorescent Aspergillus reporter conidia in conjunction with live-cell video microscopy and flow cytometry. Generated data provide insight into host cell-Aspergillus interactions such as fungicidal activity, phagocytosis, cell migration and inhibition of fungal growth.

Aspergillus fumigatus is an opportunistic fungal pathogen causing invasive infections in immunocompromised hosts with a high case-fatality rate. Research ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

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 ...

Microfluidic Tools for Probing Fungal-Microbial Interactions at the Cellular Level

Filamentous fungi are successful inhabitants of soil and play a major role in soil ecosystems, such as in the decomposition of organic and inorganic matter, as well as regulation of nutrient levels. There they also find numerous opportunities to interact with a variety of other microbes such as bacteria or other fungi. Studying fungal interactions at the cellular level, however, can be challenging owing to the black box-like nature of soil. New microfluidic tools are being developed for the study of fungal interactions; two platforms designed to study bacterial-fungal and fungal-fungal interactions are highlighted. Within these microchannels, fungal-microbial interactions can be monitored in controlled physico-chemical environments at higher temporal and spatial resolution than previously possible. Application of these tools have yielded numerous novel biological insights, such as the observation of bacterial polar attachment to hyphae or revealing uncharacterised fungal-fungal antagonisms. A key feature of these methodologies regards the ease of use of this tool by non-experts, yielding highly translatable technologies for use in microbiology labs.

Owing to the opacity of soil, interactions between its constituent microbes cannot easily be visualised with cellular resolution. Here, two microfluidic tools are presented, which offer new opportunities for investigating fungal-microbial interactions. The devices are versatile and simple to use, enabling high spatiotemporal control and high-resolution imaging at the cellular level.

Filamentous fungi are successful inhabitants of soil and play a major role in soil ecosystems, such as in the decomposition of organic and inorganic ...

Bioengineering

Split Green Fluorescent Protein System to Visualize Effectors Delivered from Bacteria During Infection

Bacteria, one of the most important causative agents of various plant diseases, secrete a set of effector proteins into the host plant cell to subvert the plant immune system. During infection cytoplasmic effectors are delivered to the host cytosol via a type III secretion system (T3SS). After delivery into the plant cell, the effector(s) targets the specific compartment(s) to modulate host cell processes for survival and replication of the pathogen. Although there has been some research on the subcellular localization of effector proteins in the host cells to understand their function in pathogenicity by using fluorescent proteins, investigation of the dynamics of effectors directly injected from bacteria has been challenging due to the incompatibility between the T3SS and fluorescent proteins.
Here, we describe our recent method of an optimized split superfolder green fluorescent protein system (sfGFPOPT) to visualize the localization of effectors delivered via the bacterial T3SS in the host cell. The sfGFP11 (11th &#946;-strand of sfGFP)-tagged effector secreted through the T3SS can be assembled with a specific organelle targeted sfGFP1-10OPT (1-10th &#946;-strand of sfGFP) leading to fluorescence emission at the site. This protocol provides a procedure to visualize the reconstituted sfGFP fluorescence signal with an effector protein from Pseudomonas syringae in a particular organelle in the Arabidopsis and Nicotiana benthamiana plants.

Fluorescent protein-based approaches to monitor effectors secreted by bacteria into host cells are challenging. This is due to the incompatibility between fluorescent proteins and the type-III secretion system. Here, an optimized split superfolder GFP system is used for visualization of effectors secreted by bacteria into the host plant cell.

Bacteria, one of the most important causative agents of various plant diseases, secrete a set of effector proteins into the host plant cell to subvert the ...

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

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

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

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

Visualization of Cellular Interactions Between the Maize Sheaths and Fungi

Detached Maize Sheaths for Live-Cell Imaging of Infection by Fungal Foliar Maize Pathogens

We have optimized a protocol to inoculate maize leaf sheaths with hemibiotrophic and necrotrophic foliar pathogenic fungi. The method is modified from one originally applied to rice leaf sheaths and allows direct microscopic observation of fungal growth and development in living plant cells. Leaf sheaths collected from maize seedlings with two fully emerged leaf collars are inoculated with 20 &#181;L drops of 5 x 105 spores/mL fungal spore suspensions and incubated in humidity chambers at 23 &#176;C under continuous fluorescent light. After 24-72 h, excess tissue is removed with a razor blade to leave a single layer of epidermal cells, an optically clear sample that can be imaged directly without the necessity for chemical fixation or clearing. Plant and fungal cells remain alive for the duration of the experiment and interactions can be visualized in real-time. Sheaths can be stained or subjected to plasmolysis to study the developmental cytology and viability of host and pathogen cells during infection and colonization. Fungal strains transformed to express fluorescent proteins can be inoculated or co-inoculated on the sheaths for increased resolution and to facilitate the evaluation of competitive or synergistic interactions. Fungal strains expressing fluorescent fusion proteins can be used to track and quantify the production and targeting of these individual proteins in planta. Inoculated sheath tissues can be extracted to characterize nucleic acids, proteins, or metabolites. The use of these sheath assays has greatly advanced the detailed studies of the mechanisms of fungal pathogenicity in maize and also of fungal protein effectors and secondary metabolites contributing to pathogenicity.

Author Spotlight: Investigating Fungal Pathogenicity Mechanisms in Maize 

This manuscript details an optimized inoculation protocol that uses detached maize leaf sheaths for reproducible cytological, physiological, and molecular studies of maize interactions with fungal plant pathogens. The leaf sheaths facilitate real-time observation of cellular interactions between the living plant and fungus in unfixed tissues.

We have optimized a protocol to inoculate maize leaf sheaths with hemibiotrophic and necrotrophic foliar pathogenic fungi. The method is modified from one ...

Real-Time Assessment of the Antifungal Activity of Primary Human Immune Cells

Source: Brunel, S. F. et al., <a href="https://app.jove.com/v/55444">Live Imaging of Antifungal Activity by Human Primary Neutrophils and Monocytes in Response to A. fumigatus.</a>J. Vis. Exp. (2017)
This video demonstrates live cell imaging for the analysis of antifungal activity in human neutrophils using fluorescent Aspergillus reporter conidia. Initially, both red and magenta fluorescence are detected within neutrophils, while red fluorescence decreases over time, indicating the degradation of conidia and confirming the antifungal activity of neutrophils.

All procedures involving human participants have been performed in compliance with the institutional, national, and international guidelines for human ...

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

Summary

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Chapters in this video

Live-cell Video Microscopy of Fungal Pathogen Phagocytosis

Live-cell Imaging and Quantitative Analysis of Embryonic Epithelial Cells in Xenopus laevis

Live Imaging of Antifungal Activity by Human Primary Neutrophils and Monocytes in Response to A. fumigatus

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

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

Microfluidic Tools for Probing Fungal-Microbial Interactions at the Cellular Level

Split Green Fluorescent Protein System to Visualize Effectors Delivered from Bacteria During Infection

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

Detached Maize Sheaths for Live-Cell Imaging of Infection by Fungal Foliar Maize Pathogens

Real-Time Assessment of the Antifungal Activity of Primary Human Immune Cells