We thank our lab members for their comments on this article. This work was funded by a start-up grant from the University of South Florida (PE).

1. General growth conditions
<ol>
	<li>Inoculate 2 mL of appropriate growth medium supplemented with antibiotics (where required) with a single colony of the strain(s) to be imaged. Incubate these seed cultures overnight at 22 &#176;C in a shaking incubator. 
	NOTE: The specific bacterial growth conditions used in this article are provided under the representative results section.</li>
	<li>Dilute overnight cultures 1:20 in fresh media in a 125 mL flask, supplement with antibiotics (where required).</li>
	<li>Grow culture(s) at 37 &#176;C in a shaking incubator until mid-logarithmic phase (OD600 = 0.5). 
	NOTE: Inducer or inhibitor could be added directly to growing culture(s) at the appropriate growth phase.</li>
	<li>Harvest cells at desired culture conditions for microscopy sample preparation as described in the following section.</li>
</ol>
2. Sample preparation
<ol>
	<li>Prepare 1% agarose by combining 0.25 g of molecular biology grade, low EEO, agarose with either 25 mL of growth medium supplemented with appropriate antibiotics (where required) for timelapse microscopy or sterile water. Heat the mixture with the help of a microwave for approximately 30 s and pour it into a sterile 100 mm x 15 mm Petri dish and let it solidify.</li>
	<li>Take a 5&#8211;50 &#181;L aliquot of the cell culture depending on the need and stain the cells. Stain cells with appropriate dyes at this stage (e.g., add fluorescent dye FM4-64 (membrane stain) for an experiment (Figure 1)).
	<ol>
		<li>Pipette a 5 &#181;L aliquot of the culture sample or stained sample to be imaged onto the bottom of a 35 mm glass bottom culture dish (with 14 mm microwell diameter and uncoated No. 1.5 coverslip; see the Table of Materials). 
		CAUTION: Do not use poly-L-lysine coated coverslips especially for timelapse microscopy as they could induce different growth pattern (we have observed this with poly-D-lysine as well; data not shown) and/or affect protein dynamics6.</li>
		<li>To minimize the use of dyes, stain 3&#8211;4 &#181;L cell suspension (harvested at OD600 = 0.5; approximately 2.7 x 107 and 3.4 x 107 for B. subtilis and S. aureus respectively per 1 mL culture) directly on the glass bottom dish.</li>
		<li>Place a pre-cut agarose slab (11 mm in diameter or of any desired size to overlap the area of the coverslip; cut using the open end of a sterile tube or a razor blade) on top of the sample and gently tap to make sure the agarose slab is lying flat against the coverslip.</li>
	</ol>
	</li>
	<li>Subsequent to imaging (see following section), if the number of cells in the field of view are not desirable, adjust the cell density of the sample via dilution or concentration by centrifugation and alter the ratio of cells in the sample using growth medium/buffer as necessary. 
	NOTE: To minimize autofluorescence emanating from the culture medium, cells can be washed in buffer (for example in standard 1x phosphate buffer saline) and resuspended in appropriate amount of buffer prior to imaging. It is possible that in this step smaller cells or vesicles may not be retained after centrifugation and will hence be lost.</li>
	<li>Add water using a pipette (~5 &#181;L drops) to the space inside the culture dish (around the coverslip) to prevent the agarose pad from drying, and to help maintain humidity during the course of image acquisition, especially for timelapse experiments.</li>
	<li>Allow culture dishes to equilibrate to the temperature inside the incubation chamber, a built-in opaque compartment provided by the manufacturer, of the microscope for 15&#8211;20 min. 
	NOTE: Turn on the heating element and set the incubation chamber to a desired temperature several hours prior to imaging. This will ensure that the hardware stabilizes to the new temperature. For prolonged timelapse imaging, place a beaker or flask with water inside the microscope chamber, away from the working area, to maintain humidity.</li>
</ol>
3. Imaging
<ol>
	<li>On the day of the experiment, turn on the microscope system. Start the imaging software (see Table of Materials) by clicking the appropriate icon on the desktop. Initialize the microscope by clicking the Initialize Microscope option (button depicted with a microscope on it) on the software&#8217;s start-up dialog box. Ensure that the objective is fully lowered using the microscope coarse adjustment prior to initialization. 
	NOTE: Following initialization three additional dialog boxes should appear in addition to the start menu: resolve3D, data collection, and filter monitor.</li>
	<li>Place a drop of 1.517 (refractive index) oil on the 100x oil immersion objective supplied by the manufacturer (Numerical Aperture = 1.4, Working Distance = 0.12 mm; see Table of Materials). 
	NOTE: It is important to choose the appropriate immersion oil for the temperature at which imaging is conducted.</li>
	<li>Load the glass bottom dish containing sample into the metal housing (coffin) and gently slide into the stage clamp.</li>
	<li>Use the coarse adjustment knob to raise the objective until the oil makes contact with the glass bottom of the dish. Use the eye piece and fine adjustment knob to bring the sample into focus. Once the cells are in focus turn the knob from the eye piece to camera mode by moving the selector switch located on the front of the microscope body to the left.</li>
	<li>Begin experiment using the imaging software. On the resolve3D window, select the Design/run experiment icon depicted by a flask. A new dialog box should appear entitled design/run experiment.
	<ol>
		<li>Set the number of Z-stacks and sample thickness using the design and then sectioning tab on the design/run experiment dialog box. 
		NOTE: For the experiments in representative results section, 17 Z-stacks at 200 nm interval for still images and four Z-stacks at 200 nm interval for timelapse microscopy were used.</li>
		<li>To measure the thickness of the cells in the sample, manually adjust the Z-plane incrementally by using the up and down arrows on the resolve3D dialog box. Mark where the cells go out-of-focus as the upper and lower limit for image acquisition. Import this information prior to running the experiment.</li>
		<li>To help minimize phototoxicity and photobleaching during timelapse imaging, reduce the number of Z-stacks and choose the mid-plane of the cells for image acquisition.</li>
	</ol>
	</li>
	<li>Select the appropriate filter set for the experiment using the design and then channels tab on the design/run experiment dialog box (TRITC: EX 542/27; EM 597/45; FITC/GFP: EX 475/28; EM 525/48; mCherry: EX 575/25; EM 632/60; Cy5: EX 632/22; EM 676/34).
	<ol>
		<li>Also, select a reference for the collection of POL/DIC information. Adjust percentage transmission (light intensity) and duration of exposure for individual channels selected prior to imaging by selecting the appropriate options on the resolve3D dialog box. 
		NOTE: In a test field of view that is not considered for the experiment test these settings to identify if the selected parameters obtain meaningful fluorescence data without missing weak signal or oversaturating it. Then import these parameters for the experimental set up.</li>
	</ol>
	</li>
	<li>Open the points list by selecting the Points list button on the resolve3D dialog box. A new dialog box should appear entitled points list. Mark several fields of view to be used in the experiment by finding appropriate fields of view using the microscope stage controls and selecting the mark point option on the points list dialog box. 
	NOTE: A replace point will have to be selected each time the microscope is refocused on a point in the points list. This can be done by focusing the microscope on the appropriate point in the list and then selecting the replace point button. It is important to not touch the analog coarse/fine adjustment knob when refocusing; use only the software to adjust the focus.</li>
	<li>Set timelapse parameters by first selecting the design and then timelapse tab on the design/run experiment dialog box. Select the timelapse check box. Enter the appropriate timelapse parameters in terms of timelapse images/total time.</li>
	<li>Set points to be imaged from the points list by first selecting the design and then points tab on the design/run experiment dialog box. Select the visit points list option and enter points to be imaged in the text box separated by commas or hyphens if it is a complete sequence.</li>
	<li>Prior to beginning an experiment, edit file names and file locations using the run tab on the design/run dialog box. File location can be changed by selecting the settings button, selecting the data folder, and by then selecting the appropriate folder or creating a new one. Change file names by entering in the new file name in the image file name text box.</li>
	<li>Begin the experiment by selecting the play button on the begin experiment dialog box. 
	NOTE: For timelapse, continually check focus at each field of view throughout the experiment using DIC setting (to avoid unnecessary photobleaching) and refocus and update the information in points list as cells tend to go out of focus over time.</li>
</ol>
4. Image processing
<ol>
	<li>Open the desired raw (R3D) image files for generation of figures. 
	NOTE: Recently saved files can be located using the data folder button on the imaging software&#8217;s start-up dialog box.</li>
	<li>Run the deconvolution program, to remove out-of-focus fluorescence light7,8, in order to produce a deconvolved D3D image file. Select the process tab on the start-up dialog box, and then select deconvolve. A new dialog box will appear entitled deconvolve.
	<ol>
		<li>Set the deconvolution parameters by dragging the appropriate image number (located in the upper left-hand corner of each image file) to the input on the deconvolution dialog box. On the deconvolution dialog box, select the more options button, and deselect the crop border rolloff after processing check box (otherwise the images cannot be superimposed over the corresponding DIC file).</li>
		<li>Click the do it button on the deconvolution dialog box to deconvolve raw image file. 
		NOTE: The deconvolution parameters could be adjusted as indicated by the software manufacturer&#8217;s guidance manual for desired results.</li>
	</ol>
	</li>
	<li>If necessary, perform manual background noise subtraction and brightness/contrast adjustment in any or all wavelength (color) channels. Do this by selecting the contrast adjustment button on the deconvolved image file.</li>
	<li>Save the image as a TIFF file by first selecting the file option at the top of the D3D dialog box. Then select save as TIFF, identify and select appropriate Z-stacks (that are within focus), select or unselect desired filter sets.</li>
	<li>Perform data quantification using any manufacturer-supplied software or freely available programs such as ImageJ5.
	<ol>
		<li>For cell size quantification click on the tool tab on the appropriate D3D image file, and then select the measure distances option. Measure distances by selecting start and end points on the desired image file using the mouse through left clicks. 
		NOTE: It is best to zoom in on the image during cell size quantification.</li>
		<li>For fluorescence signal quantification use the data inspector option under the tool tab on the appropriate D3D image file. 
		NOTE: It is best to zoom in on the image and to have only relevant filters selected when completing fluorescence signal quantification.</li>
		<li>To quantify the fluorescence signal, draw a box with column/row option set to specific dimension. Select an area with signal to be quantified and record the value. Measure the background signal by using the same size box to select an area immediately outside of the cells. Subtract the background value from the value recorded from the fluorescence signal obtained within the cell.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Microscopy has remained a mainstay in studies pertaining to microbial organisms. Given their micron-scale cell size, single-cell level studies have traditionally relied on electron microscopy (EM). Although EM has become quite a powerful technique in recent years, it has its own intrinsic limitations in addition to limited user access16. Improvements in fluorescence microscopy techniques and development of different fluorescent probes, such as FDAA3, have provided microbial...

The field of bacterial cell biology has been significantly enhanced by recent advancements in microscopy techniques1,2. Among other instruments, microscopes that are capable of conducting timelapse fluorescence microscopy experiments remain a valuable tool. Investigators can monitor various physiological events in real-time using fluorescent proteins such as, green fluorescent protein (GFP)-based transcriptional and translational reporter fusions, fluorescent D-amino acids (FDAA)3, or use other stains for labeling the cell wall, membrane and DNA. It is therefore of no surprise that fluorescence microscopy remains popular among microbial cell biologists. In addition to simply showing the end phenotypes, providing information as to how the observed phenotypes arise using timelapse microscopy could add significant value to the findings and potentially offer clues as to what cellular processes are being targeted by potential drug candidates4.
The protocols to conduct high-resolution imaging using a fully motorized, inverted, wide-field fluorescence microscope (see the Table of Materials) are provided in this article. These protocols could be adapted to suit the needs of other fluorescence microscopes that are capable of conducting timelapse microscopy. Although the software discussed here corresponds to the specific manufacturer-supplied software as indicated in the Table of Materials, software commonly supplied by other microscope manufacturers or the freely available ImageJ5, have equivalent tools for analyzing microscopy data. For conditions where timelapse is not conducive, time-course experiments could be conducted as described in this article. The protocols described here provide a detailed guide to study the phenotypic changes in two different bacterial species: B. subtilis and S. aureus. See Table 1 for strains used.

<table>
	<tbody>
		<tr>
			<td>Agarose</td>
			<td>Fisher BioReagents</td>
			<td>BP160-100</td>
			<td>Molecular Biology Grade - Low EEO</td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Invitrogen</td>
			<td>D3571</td>
			<td>Microscopy</td>
		</tr>
		<tr>
			<td>FM4-64</td>
			<td>Invitrogen</td>
			<td>T3166</td>
			<td>Microscopy</td>
		</tr>
		<tr>
			<td>Glass bottom dish</td>
			<td>MatTek</td>
			<td>P35G-1.5-14-C</td>
			<td>Microscopy</td>
		</tr>
		<tr>
			<td>IPTG</td>
			<td>Fisher BioReagents</td>
			<td>BP1755-10</td>
			<td>Dioxane-free</td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>GE</td>
			<td>DeltaVision Elite</td>
			<td>Customized Olympus IX-71 Inverted Microscope Stand; Custom Illumination Tower and Transmitted Light Illuminator Module. Objectives: PLAPON 60X (N.A. 1.42, WD 0.15 mm); OLY 100X OIL (N.A. 1.4, WD 0.12 mm); DIC Prism Nomarski for 100X Objective; CoolSnap HQ2 camera; SSI Assembly 7-color; Environmental control chamber - opaque.</td>
		</tr>
		<tr>
			<td>PC190723</td>
			<td>MilliporeSigma</td>
			<td>3445805MG</td>
			<td>FtsZ inhibitor</td>
		</tr>
		<tr>
			<td>SoftWorx</td>
			<td>GE</td>
			<td></td>
			<td>Manufacturer-supplied imaging software</td>
		</tr>
	</tbody>
</table>

live-cell fluorescence microscopy to investigate subcellular protein localization and cell morphology changes in bacteria

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

GpsB phenotypes 
Previously we have shown that Sa-GpsB is an essential protein as depletion of GpsB using an antisense RNA results in cell lysis9. Here we describe how the emergence of various cell division phenotypes and changes in protein localization could be captured using the timelapse microscopy protocol described in this article. For this purpose, S. aureus strains RB143 [SH1000 harboring pEPSA5 (empty vector)] and GGS8 [SH1000 harboring pGG59 (P...

Watch this Scientific Journal Video about Live-Cell Fluorescence Microscopy to Investigate Subcellular Protein Localization and Cell Morphology Changes in Bacteria at JoVE.com

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

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

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

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6.9K Views.  University of South Florida. This article provides a step-by-step guide to investigate protein subcellular localization dynamics and to monitor morphological changes using high-resolution fluorescence microscopy in Bacillus subtilis and Staphylococcus aureus.

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

Live Cell Imaging of Bacillus subtilis and Streptococcus pneumoniae using Automated Time-lapse Microscopy

During the last few years scientists became increasingly aware that average data obtained from microbial population based experiments are not representative of the behavior, status or phenotype of single cells. Due to this new insight the number of single cell studies rises continuously (for recent reviews see 1,2,3). However, many of the single cell techniques applied do not allow monitoring the development and behavior of one specific single cell in time (e.g. flow cytometry or standard microscopy).
Here, we provide a detailed description of a microscopy method used in several recent studies 4, 5, 6, 7, which allows following and recording (fluorescence of) individual bacterial cells of Bacillus subtilis and Streptococcus pneumoniae through growth and division for many generations. The resulting movies can be used to construct phylogenetic lineage trees by tracing back the history of a single cell within a population that originated from one common ancestor. This time-lapse fluorescence microscopy method cannot only be used to investigate growth, division and differentiation of individual cells, but also to analyze the effect of cell history and ancestry on specific cellular behavior. Furthermore, time-lapse microscopy is ideally suited to examine gene expression dynamics and protein localization during the bacterial cell cycle. The method explains how to prepare the bacterial cells and construct the microscope slide to enable the outgrowth of single cells into a microcolony. In short, single cells are spotted on a semi-solid surface consisting of growth medium supplemented with agarose on which they grow and divide under a fluorescence microscope within a temperature controlled environmental chamber. Images are captured at specific intervals and are later analyzed using the open source software ImageJ.

Live Cell Imaging of Bacillus subtilis and Streptococcus pneumoniae using Automated Time-lapse Microscopy

This protocol provides a step-by-step procedure to monitor single cell behavior of different bacteria in time using automated fluorescence time-lapse microscopy. Furthermore, we provide guidelines how to analyze the microscopy images.

During the last few years scientists became increasingly aware that average data obtained from microbial population based experiments are not ...

Visualization of Bacterial Toxin Induced Responses Using Live Cell Fluorescence Microscopy

Bacterial toxins bind to cholesterol in membranes, forming pores that allow for leakage of cellular contents and influx of materials from the external environment. The cell can either recover from this insult, which requires active membrane repair processes, or else die depending on the amount of toxin exposure and cell type1. In addition, these toxins induce strong inflammatory responses in infected hosts through activation of immune cells, including macrophages, which produce an array of pro-inflammatory cytokines2. Many Gram positive bacteria produce cholesterol binding toxins which have been shown to contribute to their virulence through largely uncharacterized mechanisms.
Morphologic changes in the plasma membrane of cells exposed to these toxins include their sequestration into cholesterol-enriched surface protrusions, which can be shed into the extracellular space, suggesting an intrinsic cellular defense mechanism3,4. This process occurs on all cells in the absence of metabolic activity, and can be visualized using EM after chemical fixation4. In immune cells such as macrophages that mediate inflammation in response to toxin exposure, induced membrane vesicles are suggested to contain cytokines of the IL-1 family and may be responsible both for shedding toxin and disseminating these pro-inflammatory cytokines5,6,7. A link between IL-1&beta; release and a specific type of cell death, termed pyroptosis has been suggested, as both are caspase-1 dependent processes8. To sort out the complexities of this macrophage response, which includes toxin binding, shedding of membrane vesicles, cytokine release, and potentially cell death, we have developed labeling techniques and fluorescence microscopy methods that allow for real time visualization of toxin-cell interactions, including measurements of dysfunction and death (Figure 1). Use of live cell imaging is necessary due to limitations in other techniques. Biochemical approaches cannot resolve effects occurring in individual cells, while flow cytometry does not offer high resolution, real-time visualization of individual cells. The methods described here can be applied to kinetic analysis of responses induced by other stimuli involving complex phenotypic changes in cells.

Methods for purifying the cholesterol binding toxin streptolysin O from recombinant E. coli and visualization of toxin binding to live eukaryotic cells are described. Localized delivery of toxin induces rapid and complex changes in targeted cells revealing novel aspects of toxin biology.

Bacterial toxins bind to cholesterol in membranes, forming pores that allow for leakage of cellular contents and influx of materials from the external ...

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 ...

Fluorescence Microscopy Methods for Determining the Viability of Bacteria in Association with Mammalian Cells

Central to the field of bacterial pathogenesis is the ability to define if and how microbes survive after exposure to eukaryotic cells. Current protocols to address these questions include colony count assays, gentamicin protection assays, and electron microscopy. Colony count and gentamicin protection assays only assess the viability of the entire bacterial population and are unable to determine individual bacterial viability. Electron microscopy can be used to determine the viability of individual bacteria and provide information regarding their localization in host cells. However, bacteria often display a range of electron densities, making assessment of viability difficult. This article outlines protocols for the use of fluorescent dyes that reveal the viability of individual bacteria inside and associated with host cells. These assays were developed originally to assess survival of Neisseria gonorrhoeae in primary human neutrophils, but should be applicable to any bacterium-host cell interaction. These protocols combine membrane-permeable fluorescent dyes (SYTO9 and 4&#39;,6-diamidino-2-phenylindole [DAPI]), which stain all bacteria, with membrane-impermeable fluorescent dyes (propidium iodide and SYTOX Green), which are only accessible to nonviable bacteria. Prior to eukaryotic cell permeabilization, an antibody or fluorescent reagent is added to identify extracellular bacteria. Thus these assays discriminate the viability of bacteria adherent to and inside eukaryotic cells. A protocol is also provided for using the viability dyes in combination with fluorescent antibodies to eukaryotic cell markers, in order to determine the subcellular localization of individual bacteria. The bacterial viability dyes discussed in this article are a sensitive complement and/or alternative to traditional microbiology techniques to evaluate the viability of individual bacteria and provide information regarding where bacteria survive in host cells.

Central to the field of bacterial pathogenesis is the ability to define if and how microbes survive after exposure to eukaryotic cells. This article outlines protocols for the use of fluorescent dyes that reveal the viability of individual bacteria inside and associated with host cells.

Central to the field of bacterial pathogenesis is the ability to define if and how microbes survive after exposure to eukaryotic cells. Current protocols ...

Fluorescence in situ Hybridizations (FISH) for the Localization of Viruses and Endosymbiotic Bacteria in Plant and Insect Tissues

Fluorescence in situ hybridization (FISH) is a name given to a variety of techniques commonly used for visualizing gene transcripts in eukaryotic cells and can be further modified to visualize other components in the cell such as infection with viruses and bacteria. Spatial localization and visualization of viruses and bacteria during the infection process is an essential step that complements expression profiling experiments such as microarrays and RNAseq in response to different stimuli. Understanding the spatiotemporal infections with these agents complements biological experiments aimed at understanding their interaction with cellular components. Several techniques for visualizing viruses and bacteria such as reporter gene systems or immunohistochemical methods are time-consuming, and some are limited to work with model organisms&#160;and involve complex methodologies. FISH that targets RNA or DNA species in the cell is a relatively easy and fast method for studying spatiotemporal localization of genes and for diagnostic purposes. This method can be robust and relatively easy to implement when the protocols employ short hybridizing, commercially-purchased probes, which are not expensive. This is particularly robust when sample preparation, fixation, hybridization, and microscopic visualization do not involve complex steps. Here we describe a protocol for localization of bacteria and viruses in insect and plant tissues. The method is based on simple preparation, fixation, and hybridization of insect whole mounts and dissected organs or hand-made plant sections, with 20 base pairs short DNA probes conjugated to fluorescent dyes on their 5&#39; or 3&#39; ends. This protocol has been successfully applied to a number of insect and plant tissues, and can be used to analyze expression of mRNAs or other RNA or DNA species in the cell.

Fluorescence in situ Hybridizations FISH for the Localization of Viruses and Endosymbiotic Bacteria in Plant and Insect Tissues

We describe here a simple fluorescence in situ hybridization (FISH) method for the localization of viruses and bacteria in insect and plant tissues. This protocol can be extended for the visualization of mRNA in whole mount and microscopic sections.

Fluorescence in situ hybridization (FISH) is a name given to a variety of techniques commonly used for visualizing gene transcripts in eukaryotic cells ...

Identification of Plasmodesmal Localization Sequences in Proteins In Planta

Plasmodesmata (Pd) are cell-to-cell connections that function as gateways through which small and large molecules are transported between plant cells. Whereas Pd transport of small molecules, such as ions and water, is presumed to occur passively, cell-to-cell transport of biological macromolecules, such proteins, most likely occurs via an active mechanism that involves specific targeting signals on the transported molecule. The scarcity of identified plasmodesmata (Pd) localization signals (PLSs) has severely restricted the understanding of protein-sorting pathways involved in plant cell-to-cell macromolecular transport and communication. From a wealth of plant endogenous and viral proteins known to traffic through Pd, only three PLSs have been reported to date, all of them from endogenous plant proteins. Thus, it is important to develop a reliable and systematic experimental strategy to identify a functional PLS sequence, that is both necessary and sufficient for Pd targeting, directly in the living plant cells. Here, we describe one such strategy using as a paradigm the cell-to-cell movement protein (MP) of the Tobacco mosaic virus (TMV). These experiments, that identified and characterized the first plant viral PLS, can be adapted for discovery of PLS sequences in most Pd-targeted proteins.

Identification of Plasmodesmal Localization Sequences in Proteins In Planta

Plant intercellular connections, the plasmodesmata (Pd), play central roles in plant physiology and plant-virus interactions. Critical to Pd transport are sorting signals that direct proteins to Pd. However, our knowledge about these sequences is still in its infancy. We describe a strategy to identify Pd localization signals in Pd-targeted proteins.

Plasmodesmata (Pd) are cell-to-cell connections that function as gateways through which small and large molecules are transported between plant cells. ...

In Vitro Canine Neutrophil Extracellular Trap Formation: Dynamic and Quantitative Analysis by Fluorescence Microscopy

In response to invading pathogens, neutrophils release neutrophil extracellular traps (NETs), which are extracellular networks of DNA decorated with histones and antimicrobial proteins. Excessive NET formation (NETosis) and citH3 release during sepsis is associated with multiple organ dysfunction and mortality in mice and humans but its implications in dogs are unknown. Herein, we describe a technique to isolate canine neutrophils from whole blood for observation and quantification of NETosis. Leukocyte-rich plasma, generated by dextran sedimentation, is separated by commercially available density gradient separation media and granulocytes collected for cell count and viability testing. To observe real-time NETosis in live neutrophils, cell permeant and cell impermeant fluorescent nucleic acid stains are added to neutrophils activated either by lipopolysaccharide (LPS) or phorbol 12-myristate 13-acetate (PMA). Changes in nuclear morphology and NET formation are observed over time by fluorescence microscopy. In vitro NETosis is further characterized by co-colocalization of cell-free DNA (cfDNA), myeloperoxidase (MPO) and citrullinated histone H3 (citH3) using a modified double-immunolabelling protocol. To objectively quantify NET formation and citH3 expression using fluorescence microscopy, NETs and citH3-positive cells are quantified in a blinded manner using available software. This technique is a specific assay to evaluate the in vitro capacity of canine neutrophils to undergo NETosis.

In Vitro Canine Neutrophil Extracellular Trap Formation: Dynamic and Quantitative Analysis by Fluorescence Microscopy

We describe methods to isolate canine neutrophils from whole blood and visualize NET formation in live neutrophils using fluorescence microscopy. Also described are protocols to quantify NET formation and citrullinated histone H3 (citH3) expression using immunofluorescence microscopy.

In response to invading pathogens, neutrophils release neutrophil extracellular traps (NETs), which are extracellular networks of DNA decorated with ...

Quantification of Efferocytosis by Single-cell Fluorescence Microscopy

Studying the regulation of efferocytosis requires methods that are able to accurately quantify the uptake of apoptotic cells and to probe the signaling and cellular processes that control efferocytosis. This quantification can be difficult to perform as apoptotic cells are often efferocytosed piecemeal, thus necessitating methods which can accurately delineate between the efferocytosed portion of an apoptotic target versus residual unengulfed cellular fragments. The approach outlined herein utilizes dual-labeling approaches to accurately quantify the dynamics of efferocytosis and efferocytic capacity of efferocytes such as macrophages. The cytosol of the apoptotic cell is labeled with a cell-tracking dye to enable monitoring of all apoptotic cell-derived materials, while surface biotinylation of the apoptotic cell allows for differentiation between internalized and non-internalized apoptotic cell fractions. The efferocytic capacity of efferocytes is determined by taking fluorescent images of live or fixed cells and quantifying the amount of bound versus internalized targets, as differentiated by streptavidin staining. This approach offers several advantages over methods such as flow cytometry, namely the accurate delineation of non-efferocytosed versus efferocytosed apoptotic cell fractions, the ability to measure efferocytic dynamics by live-cell microscopy, and the capacity to perform studies of cellular signaling in cells expressing fluorescently-labeled transgenes. Combined, the methods outlined in this protocol serve as the basis for a flexible experimental approach that can be used to accurately quantify efferocytic activity and interrogate cellular signaling pathways active during efferocytosis.

Efferocytosis, the phagocytic removal of apoptotic cells, is required to maintain homeostasis and is facilitated by receptors and signaling pathways that allow for the recognition, engulfment, and internalization of apoptotic cells. Herein, we present a fluorescence microscopy protocol for the quantification of efferocytosis and the activity of efferocytic signaling pathways.

Studying the regulation of efferocytosis requires methods that are able to accurately quantify the uptake of apoptotic cells and to probe the signaling ...

Live-Cell Forward Genetic Approach to Identify and Isolate Developmental Mutants in Chlamydia trachomatis

The intracellular bacterial pathogen Chlamydia trachomatis undergoes a developmental cycle consisting of two morphologically discrete developmental forms. The non-replicative elementary body (EB) initiates infection of the host. Once inside, the EB differentiates into the reticulate body (RB). The RB then undergoes multiple rounds of replication, before differentiating back to the infectious EB form. This cycle is essential for chlamydial survival as failure to switch between cell types prevents either host invasion or replication.
Limitations in genetic techniques due to the obligate intracellular nature of Chlamydia have hampered identification of the molecular mechanisms involved in the cell-type development. We designed a novel dual promoter-reporter plasmid system that, in conjunction with live-cell microscopy, allows for the visualization of cell type switching in real time. To identify genes involved in the regulation of cell-type development, the live-cell promoter-reporter system was leveraged for the development of a forward genetic approach by combining chemical mutagenesis of the dual reporter strain, imaging and tracking of Chlamydia with altered developmental kinetics, followed by clonal isolation of mutants. This forward genetic workflow is a flexible tool that can be modified for directed interrogation into a wide range of genetic pathways.

Live-Cell Forward Genetic Approach to Identify and Isolate Developmental Mutants in Chlamydia trachomatis

This protocol utilizes fluorescent promoter-reporters, live-cell microscopy, and individual inclusion extraction in a directed forward genetic approach to identify and isolate developmental mutants of Chlamydia trachomatis.

The intracellular bacterial pathogen Chlamydia trachomatis undergoes a developmental cycle consisting of two morphologically discrete developmental forms. ...

Intravital Widefield Fluorescence Microscopy of Pulmonary Microcirculation in Experimental Acute Lung Injury Using a Vacuum-Stabilized Imaging System

Intravital imaging of leukocyte-endothelial interactions offers valuable insights into immune-mediated disease in live animals. The study of acute lung injury (ALI)/acute respiratory distress syndrome (ARDS) and other respiratory pathologies in vivo is difficult due to the limited accessibility and inherent motion artifacts of the lungs. Nonetheless, various approaches have been developed to overcome these challenges. This protocol describes a method for intravital fluorescence microscopy to study real-time leukocyte-endothelial interactions in the pulmonary microcirculation in an experimental model of ALI. An in vivo lung imaging system and 3-D printed intravital microscopy platform are used to secure the anesthetized mouse and stabilize the lung while minimizing confounding lung injury. Following preparation, widefield fluorescence microscopy is used to study leukocyte adhesion, leukocyte rolling, and capillary function. While the protocol presented here focuses on imaging in an acute model of inflammatory lung disease, it may also be adapted to study other pathological and physiological processes in the lung.

Intravital fluorescence microscopy can be utilized to study leukocyte-endothelial interactions and capillary perfusion in real-time. This protocol describes methods to image and quantify these parameters in the pulmonary microcirculation using a vacuum-stabilized lung imaging system.

Intravital imaging of leukocyte-endothelial interactions offers valuable insights into immune-mediated disease in live animals. The study of acute lung ...