We thank Asya Smirnov and Laura Gonyar for critical reading of the manuscript. This work was supported by grants NIH R00 TW008042 and R01 AI097312 to A.K.C. M.B.J. was supported in part by NIH T32 AI007046.

1. Assessing Bacterial Viability with Propidium Iodide and SYTO9
<ol>
	<li>Infect cells that are adherent to 12 mm diameter circular glass coverslips in 24-well plates with bacteria of interest. Do NOT fix the cells with aldehydes or organic solvents.</li>
	<li>Rinse cells once gently in 0.1 M 3-(N-morpholino)propanesulfonic acid (MOPS), pH 7.2, containing 1 mM MgCl2 (MOPS/MgCl2).</li>
	<li>Incubate cells for 10 min in the dark at room temperature with Alexa Fluor 647-coupled antibody or lectin that binds to the bacterial species of interest, in MOPS/MgCl2, to detect external bacteria.</li>
</ol>
NOTE: Conduct controls with live and dead bacteria, in the absence of host cells, to show that the lectin or antibody of interest binds all bacteria regardless of viability (Figure 1).
<ol start="4">
	<li>Rinse cells two times with MOPS/MgCl2.</li>
	<li>Aspirate media from cells and add 0.5 ml Live/Dead Staining Solution. Live/Dead Staining Solution is 5 &#956;M SYTO9, 30 &#956;M propidium iodide, and 0.1% saponin (final concentrations) in MOPS/MgCl2.</li>
	<li>Incubate cells for 15 min at room temperature in the dark.</li>
	<li>Rinse cells two times in MOPS/MgCl2.</li>
	<li>Invert coverslips face down onto glass slides and seal with clear nail polish. Do not use mounting media.</li>
	<li>Acquire images within 30 min, using a fluorescence microscope with filter sets compatible with green, red, and far-red image acquisition.</li>
</ol>
NOTE: Both conventional fluorescence microscopy and confocal laser scanning microscopy can be used. After 30 min the fluorescent dyes begin to leak from the bacteria and the data acquired are no longer accurate. Images shown in this article were acquired on a Nikon Eclipse E800 upright fluorescence microscope with Hamamatsu Orca-ER digital camera using Openlab software. Fluorescence of Alexa Fluor 647 was detected using a filter with excitation wavelength of 590 - 650 nm and an emission filter of 663 - 735 nm, and is false-colored blue. Fluorescence from propidium iodide was detected using a filter with excitation wavelength of 540 - 580 nm and an emission filter of 600 - 660 nm. Fluorescence from SYTO9 was detected using a filter with excitation wavelength of 465 - 495 nm and an emission filter of 515 - 555 nm.
<ol start="10">
	<li>This protocol will result in images where external nonviable bacteria appear blue + red, internal nonviable bacteria appear red only, external viable bacteria appear blue + green, and internal viable bacteria appear green only. Count by eye the numbers of bacteria that are external nonviable, internal nonviable, external viable, and internal viable.</li>
	<li>Calculate the percent of external viable bacteria by dividing the number of external viable bacteria by the total number of external bacteria (viable plus nonviable). Calculate the percent of internal viable bacteria by dividing the number of internal viable bacteria by the total number of internal bacteria (viable plus nonviable) (Figure 4).</li>
	<li>There are two essential controls to perform with this protocol. First, validate that all nonviable bacteria are propidium iodide-positive and all live bacteria are SYTO9-positive. A mid-logarithmic culture of bacteria (in the absence of any host cells) should be &#62;95% SYTO9-positive. Shown in Figure 2 is a mid-logarithmic culture of N. gonorrhoeae (at 108 colony forming units per ml) incubated with Live/Dead Staining Solution. Second, validate that both propidium iodide and SYTO9 can enter permeabilized, infected host cells.
	<ol>
		<li>To generate a population of dead bacteria, collect 2 x 108 bacterial colony forming units in a 1.5 ml microfuge tube, add 70% isopropanol and sit for 10 min. Pellet the bacteria in a microfuge and wash the bacteria twice in PBS to remove residual isopropanol. Then follow protocol steps 1.5 - 1.7. Add 5 &#956;l of bacterial suspension to a glass slide and overlay with a coverslip. Image samples as in protocol step 1.9. Under these conditions, 100% of the population should be propidium iodide-positive (Figure 2). In some bacterial species, propidium iodide may not completely overwhelm SYTO9 staining, and nonviable bacteria may appear yellow or orange.</li>
		<li>For phagocytic cells like neutrophils, expose the cells to isopropanol-killed bacteria and ensure that 100% of the intracellular bacteria are propidium iodide-positive (Figure 3). For nonphagocytic cells that may not internalize dead bacteria, it may be sufficient to treat infected cells with sodium azide or other cell-permeant antimicrobial agents prior to adding Live/Dead Staining Solution.</li>
	</ol>
	</li>
</ol>
2. Assessing Bacterial Viability with SYTOX Green and DAPI
<ol>
	<li>Label bacteria of interest with 10 &#956;g/ml DAPI in Morse&#39;s Defined Medium 25 for 20 min at room temperature in the dark.</li>
	<li>Infect cells that are adherent to 12 mm diameter circular glass coverslips in 24-well plates with DAPI-labeled bacteria. Do NOT fix the cells with aldehydes or organic solvents.</li>
	<li>Rinse cells once with MOPS/MgCl2.</li>
	<li>Incubate cells for 10 min at room temperature in the dark with Alexa Fluor 647-coupled antibody or lectin that binds to the bacterial species of interest, in MOPS/MgCl2, to detect external bacteria. See note in protocol step 1.3 for suggested controls.</li>
	<li>Aspirate media from cells and add 0.5 ml 0.4 &#956;M SYTOX Green in MOPS/MgCl2.</li>
	<li>Incubate cells for 5 min at room temperature in the dark.</li>
	<li>Rinse cells two times in MOPS/MgCl2.</li>
	<li>Wash cells one time in MOPS/MgCl2 for 5 min.</li>
	<li>Acquire images within 30 min with a fluorescence microscope. See protocol step 1.9 for description of microscope, digital camera, and acquisition software.</li>
</ol>
NOTE: Fluorescence from Alexa Fluor 647 was detected using a filter with excitation wavelength of 590 - 650 nm and an emission filter of 663 - 735 nm, and is false-colored red. Fluorescence from SYTOX Green was detected using a filter with excitation wavelength of 465 - 495 nm and an emission filter of 515 - 555 nm. Fluorescence from DAPI was detected using a filter with excitation wavelength of 355 - 375 nm and barrier filter of 400 nm.
<ol start="10">
	<li>This protocol will result in images where external nonviable bacteria appear red + green, internal nonviable bacteria appear green + blue, external viable bacteria appear red + blue, and internal viable bacteria appear blue only (Figure 4). Quantify the percent of external and internal bacteria as described in protocol step 1.11.</li>
	<li>The controls described in protocol step 1.12 should be performed with the DAPI/SYTOX Green dye combination (Figures 2 and 3).</li>
</ol>
3. Assessing Bacterial Viability Alongside Subcellular Localization
<ol>
	<li>Label bacteria of interest with 10 &#956;g/ml DAPI in Morse&#39;s Defined Medium for 20 min at room temperature in the dark.</li>
	<li>Infect cells that are adherent to 12 mm diameter circular glass coverslips in 24-well plates with DAPI-labeled bacteria. Do NOT fix the cells with aldehydes or organic solvents.</li>
	<li>Rinse cells once with MOPS/MgCl2.</li>
	<li>Incubate 10 min at room temperature in the dark with Alexa Fluor 647-coupled antibody or lectin that binds to the bacterial species of interest, in MOPS/MgCl2, to detect external bacteria. See note in protocol step 1.3 for suggested controls.</li>
	<li>Aspirate media from cells and rinse cells two times in MOPS/ MgCl2.</li>
	<li>Incubate cells with Alexa Fluor 555-coupled antibody against subcellular marker of interest for 20 min, in MOPS/MgCl2 containing 0.2% saponin.</li>
	<li>Rinse cells two times in MOPS/MgCl2.</li>
	<li>Wash cells one time in MOPS/MgCl2 for 5 min.</li>
	<li>Aspirate media from cells and add 0.4 &#956;M SYTOX Green in MOPS/MgCl2.</li>
	<li>Incubate cells 5 min at room temperature in the dark.</li>
	<li>Rinse cells two times in MOPS/MgCl2.</li>
	<li>Wash cells one time in MOPS/ MgCl2 for 5 min.</li>
	<li>Acquire images of slides within 30 min on fluorescence microscope. See protocol step 1.9 for description of microscope, digital camera, and acquisition software.</li>
</ol>
NOTE: Fluorescence from Alexa Fluor 647 was detected using a filter with excitation wavelength of 590 - 650 nm and emission filter of 663 - 735 nm, and is false-colored purple. Fluorescence from Alexa Fluor 555 was detected using a filter with excitation wavelength of 540 - 580 nm and an emission filter of 600 - 660 nm. Fluorescence from SYTOX Green was detected using a filter with excitation wavelength of 465 - 495 nm and an emission filter of 515 - 555 nm. Fluorescence from DAPI was detected using a filter with excitation wavelength of 355 - 375 nm and barrier filter of 400 nm.
<ol start="14">
	<li>This protocol will result in images where external nonviable bacteria appear purple + green, internal nonviable bacteria appear green + blue, external viable bacteria appear purple + blue, internal viable bacteria appear blue only, and subcellular protein appears red (Figure 4). Quantify the perecent of external and internal viable bacteria as described in protocol steps 1.11.</li>
	<li>The controls described in protocol step 1.12 should be performed with the DAPI/SYTOX Green dye combination (Figures 2 and 3).</li>
	<li>In addition to counting viable and nonviable bacteria, classify each bacteria as either positive or negative for colocalization with the subcellular marker of interest.</li>
	<li>Calculate the percent of viable bacteria colocalized with the subcellular marker by dividing the number of colocalized viable bacteria by the total number of viable bacteria. Calculate the percent of nonviable bacteria colocalized with the subcellular marker by dividing the number of nonviable colocalized bacteria by the total number of nonviable bacteria.</li>
</ol>

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The authors declare that they have no competing financial interests.

Presented here are two protocols that use DNA binding and viability dyes in conjunction with a fluorescent lectin to identify live and dead bacteria attached to and inside human cells. Since both protocols effectively discriminate live from dead bacteria, the choice of which protocol to use depends upon the goal of the experiment. The first protocol uses propidium iodide to detect nonviable bacteria and SYTO9 to detect intact bacteria. Shown in Figure 4A is a representative image of protocol 1 applied to...

There is a dynamic interaction and co-evolution between bacteria and the hosts in which they reside. Bacteria have evolved adherence organelles, secretion systems, and/or the ability to produce toxins that enable their productive infection of host phagocytic and non-phagocytic cells. The bacteria must also contend with recognition and antimicrobial activities of the host immune system. The host immune system is comprised of innate and adaptive components including physical and chemical barriers, immune cells, the complement system, and other components of humoral immunity. While many bacteria are susceptible to killing and clearance by the multilayered host immune response, some pathogenic and opportunistic bacteria have evolved mechanisms to infect a variety of host cells and subvert clearance by the host immune response 1. Neisseria gonorrhoeae is one example of a bacterial pathogen that is highly adapted to persist in its human host. N. gonorrhoeae readily colonizes the luminal surfaces of mucosal epithelial cells of the urogenital tract, pharynx, conjunctiva, and rectum. Colonization triggers the abundant recruitment of neutrophils at mucosal sites. Neutrophils are professional phagocytes that possess a variety of antimicrobial processes to kill microorganisms; however, N. gonorrhoeae is capable of surviving in the presence of neutrophils 2-5. Understanding how bacterial pathogens such as N. gonorrhoeae subvert, suppress, and hijack the immune response to ultimately survive in normally hostile host environments is crucial to the development of new therapies for combating infectious diseases.
Experimental protocols often used to investigate bacterial survival in host cells include colony count assays, gentamicin protection assays, and electron microscopy. In colony count assays, a population of infected cells is lysed (for instance, with a detergent to which the bacteria are resistant) to liberate the bacteria. The lysates are diluted and plated on agar-based media, and colony-forming units in the lysates are enumerated for each time point and/or experimental condition. This approach reports the viability of the entire bacterial population but is not capable of differentiating between intracellular and extracellular survival. A variation on the colony count assay, the gentamicin protection assay, specifically measures intracellular bacterial survival, based on the inability of the antibiotic gentamicin to cross the eukaryotic plasma membrane 6. However, this assay is dependent on the bacteria being susceptible to killing by gentamicin (or another antibiotic that is similarly eukaryotic membrane-impermeant) and the inability of the antibiotic to have access to internal bacteria. Therefore, a gentamicin protection assay may not be effective for examination of all bacterial species or when examining bacterial survival in highly pinocytic cells such as neutrophils. Neither of these approaches reveals the subcellular localization or other behavior of individual bacteria (e.g. if the bacteria form aggregates or microcolonies that behave differently from individual bacterial cells). Another frequently used approach to examine the viability of individual external and internal bacteria is thin-section transmission electron microscopy (TEM). This approach is advantageous in that it can provide information regarding the location of the bacteria in host cells (e.g. phagosome, cytoplasm, autophagosome), which can be further investigated by immunoelectron microscopy with gold-coupled antibodies against subcellular markers. However, electron microscopy is not especially sensitive at assessing bacterial viability. When embedded sections are stained with uranyl acetate, lead citrate, or other electron-dense reagents and imaged by electron microscopy, electron-dense bacteria are considered viable and electron-lucent nonviable 7,8. However, this assumption overestimates bacterial viability, since only those dead bacteria with severely disrupted membranes and devoid of cytoplasm appear electron-lucent. In addition, some bacterial species may display a range of electron densities depending on their stage of growth, making it difficult to determine viability.
As an alternative or in addition to these widely used methods, here we provide protocols and rationale for the use of fluorescent dyes that indicate bacterial viability to assess the survival of bacteria attached to and internalized by host cells. To identify extracellular bacteria, infected cells are first exposed to a fluorescent reagent, such as a lectin or bacteria-specific antibody. The infected cells are then permeabilized and exposed to DNA-specific dyes that are differentially accessible to bacteria with intact vs. degraded membranes, as a surrogate for bacterial viability. In the first protocol, the membrane permeable dye SYTO9 identifies the total bacterial population, while propidium iodide is only accessible to those bacteria that have compromised membranes and are thus considered nonviable. Propidium iodide and SYTO9 have been used to evaluate bacterial viability in biofilms, discriminate pathogenic from nonpathogenic bacteria, and enumerate viable water-borne bacteria 9-12. In the second protocol, 4&#39;,6&#39;-diamidino-2-phenylindole (DAPI) identifies total bacteria, while SYTOX Green is only accessible to the nonviable population. These viability dye pairs can be combined with immunofluorescence to determine each bacterium&#39;s location in relation to a protein of interest, for instance to define bacterial subcellular localization. The use of these assays provides key insight into the interactions that result in bacterial killing or survival during infection of host cells. The protocols outlined in this article were used to assess the viability of N. gonorrhoeae that is attached to and inside primary human neutrophils, including in different populations of neutrophil phagosomes 5,13,14. However, these protocols can be applied to assess viability of gram-positive and gram-negative bacteria in professional phagocytes, non-professional phagocytes, and protozoa 15-24.

<table border="1">
	<tbody>
		<tr>
			<td>Name of Reagent/Material</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>21 G 3/4 butterfly needles for blood collection</td>
			<td>Becton Dickinson</td>
			<td>367251</td>
			<td></td>
		</tr>
		<tr>
			<td>Blood collection tubes with Sodium Heparin 10 ml</td>
			<td>Becton Dickinson</td>
			<td>366480</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile water for irrigation</td>
			<td>Baxter</td>
			<td>07-09-64-070</td>
			<td></td>
		</tr>
		<tr>
			<td>Dextran 500</td>
			<td>Sigma</td>
			<td>31392</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Chloride</td>
			<td>Fisher Scientific</td>
			<td>S641</td>
			<td></td>
		</tr>
		<tr>
			<td>Dextrose</td>
			<td>Ricca Chemical Company</td>
			<td>RDCD0200</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s PBS no Ca2+ or Mg2+</td>
			<td>Thermo Scientific</td>
			<td>SH3002802</td>
			<td></td>
		</tr>
		<tr>
			<td>Ficoll solution</td>
			<td>GE Healthcare</td>
			<td>17-1440-03</td>
			<td></td>
		</tr>
		<tr>
			<td>Acetic Acid</td>
			<td>Fisher Scientific</td>
			<td>BP2401</td>
			<td></td>
		</tr>
		<tr>
			<td>12 mm circular glass coverslips</td>
			<td>Fisher Scientific</td>
			<td>12-545-80 12CIR-1</td>
			<td></td>
		</tr>
		<tr>
			<td>24-well plates</td>
			<td>Corning Incorporated</td>
			<td>3524</td>
			<td></td>
		</tr>
		<tr>
			<td>Pooled Human Serum</td>
			<td>Sigma</td>
			<td>S7023</td>
			<td></td>
		</tr>
		<tr>
			<td>RPMI</td>
			<td>Mediatech</td>
			<td>15-040-CV</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>Thermo Scientific</td>
			<td>SH3007103</td>
			<td></td>
		</tr>
		<tr>
			<td>Human interleukin-8</td>
			<td>R&#38;D Systems</td>
			<td>208-IL/CF</td>
			<td></td>
		</tr>
		<tr>
			<td>MOPS</td>
			<td>Sigma</td>
			<td>M3183</td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>Fisher Scientific</td>
			<td>BP214</td>
			<td></td>
		</tr>
		<tr>
			<td>Propidium Iodide</td>
			<td>Life Technologies</td>
			<td>L7007 or L7012</td>
			<td></td>
		</tr>
		<tr>
			<td>SYTO9</td>
			<td>Life Technologies</td>
			<td>L7007 or L7012</td>
			<td></td>
		</tr>
		<tr>
			<td>Saponin</td>
			<td>Fluka Analytical</td>
			<td>47036</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor 647-coupled soybean lectin</td>
			<td>Life Technologies</td>
			<td>L-32463</td>
			<td></td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Sigma</td>
			<td>D8417</td>
			<td></td>
		</tr>
		<tr>
			<td>SYTOX Green</td>
			<td>Life Technologies</td>
			<td>S7020</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse anti-CD63</td>
			<td>Developmental Studies Hybridoma Bank</td>
			<td>H5C6</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor 555 Antibody Labeling Kit</td>
			<td>Life Technologies</td>
			<td>A20187</td>
			<td></td>
		</tr>
		<tr>
			<td>Hemacytometer Bright Line</td>
			<td>Hausser Scientific</td>
			<td>1492</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>EMS</td>
			<td>78320</td>
			<td></td>
		</tr>
		<tr>
			<td>Sorvall Legend RT + Centrifuge</td>
			<td>Thermo Scientific</td>
			<td>75004377</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

The protocols outlined were used to examine survival of N. gonorrhoeae after exposure to primary human neutrophils 5,26. Neutrophils were infected with N. gonorrhoeae and processed with protocol 1, using the green-fluorescent viability dye SYTO9 and the red-fluorescent propidium iodide (Figure 4A). The dyes were added in the presence of saponin, which sequesters cholesterol to preferentially permeabilize host cell plasma membranes, not N. gonorrhoeae membranes. Other...

Watch this Scientific Journal Video about Fluorescence Microscopy Methods for Determining the Viability of Bacteria in Association with Mammalian Cells at JoVE.com

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. 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-microscopy-methods-for-determining-viability-bacteria

Research

JoVE Journal

Immunology and Infection

43.5K Views.  University of Virginia Health Sciences Center. 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.

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

Following Cell-fate in E. coli After Infection by Phage Lambda

The system comprising bacteriophage (phage) lambda and the bacterium E. coli has long served as a paradigm for cell-fate determination1,2. Following the simultaneous infection of the cell by a number of phages, one of two pathways is chosen: lytic (virulent) or lysogenic (dormant)3,4. We recently developed a method for fluorescently labeling individual phages, and were able to examine the post-infection decision in real-time under the microscope, at the level of individual phages and cells5. Here, we describe the full procedure for performing the infection experiments described in our earlier work5. This includes the creation of fluorescent phages, infection of the cells, imaging under the microscope and data analysis. The fluorescent phage is a "hybrid", co-expressing wild- type and YFP-fusion versions of the capsid gpD protein. A crude phage lysate is first obtained by inducing a lysogen of the gpD-EYFP (Enhanced Yellow Fluorescent Protein) phage, harboring a plasmid expressing wild type gpD. A series of purification steps are then performed, followed by DAPI-labeling and imaging under the microscope. This is done in order to verify the uniformity, DNA packaging efficiency, fluorescence signal and structural stability of the phage stock. The initial adsorption of phages to bacteria is performed on ice, then followed by a short incubation at 35&deg;C to trigger viral DNA injection6. The phage/bacteria mixture is then moved to the surface of a thin nutrient agar slab, covered with a coverslip and imaged under an epifluorescence microscope. The post-infection process is followed for 4 hr, at 10 min interval. Multiple stage positions are tracked such that ~100 cell infections can be traced in a single experiment. At each position and time point, images are acquired in the phase-contrast and red and green fluorescent channels. The phase-contrast image is used later for automated cell recognition while the fluorescent channels are used to characterize the infection outcome: production of new fluorescent phages (green) followed by cell lysis, or expression of lysogeny factors (red) followed by resumed cell growth and division. The acquired time-lapse movies are processed using a combination of manual and automated methods. Data analysis results in the identification of infection parameters for each infection event (e.g. number and positions of infecting phages) as well as infection outcome (lysis/lysogeny). Additional parameters can be extracted if desired.

Following Cell-fate in E. coli After Infection by Phage Lambda

This article describes the procedure for preparing a fluorescently-labeled version of bacteriophage lambda, infection of E. coli bacteria, following the infection outcome under the microscope, and analysis of infection results.

The system comprising bacteriophage (phage) lambda and the bacterium E. coli has long served as a paradigm for cell-fate determination1,2. Following the ...

Tractable Mammalian Cell Infections with Protozoan-primed Bacteria

Many intracellular bacterial pathogens use freshwater protozoans as a natural reservoir for proliferation in the environment. Legionella pneumophila, the causative agent of Legionnaires' pneumonia, gains a pathogenic advantage over in vitro cultured bacteria when first harvested from protozoan cells prior to infection of mammalian macrophages. This suggests that important virulence factors may not be properly expressed in vitro. We have developed a tractable system for priming L. pneumophila through its natural protozoan host Acanthamoeba castellanii prior to mammalian cell infection. The contribution of any virulence factor can be examined by comparing intracellular growth of a mutant strain to wild-type bacteria after protozoan priming. GFP-expressing wild-type and mutant L. pneumophila strains are used to infect protozoan monolayers in a priming step and allowed to reach late stages of intracellular growth. Fluorescent bacteria are then harvested from these infected cells and normalized by spectrophotometry to generate comparable numbers of bacteria for a subsequent infection into mammalian macrophages. For quantification, live bacteria are monitored after infection using fluorescence microscopy, flow cytometry, and by colony plating. This technique highlights and relies on the contribution of host cell-dependent gene expression by mimicking the environment that would be encountered in a natural acquisition route. This approach can be modified to accommodate any bacterium that uses an intermediary host as a means for gaining a pathogenic advantage.

This technique provides a method to harvest, normalize and quantify intracellular growth of bacterial pathogens that are pre-cultivated in natural protozoan host cells prior to infections of mammalian cells. This method can be modified to accommodate a wide variety of host cells for the priming stage as well as target cell types.

Many intracellular bacterial pathogens use freshwater protozoans as a natural reservoir for proliferation in the environment. Legionella pneumophila, the ...

Use of Image Cytometry for Quantification of Pathogenic Fungi in Association with Host Cells

Studies of the cellular pathogenesis mechanisms of pathogenic yeasts such as Candida albicans, Histoplasma capsulatum, and Cryptococcus neoformans commonly employ infection of mammalian hosts or host cells (i.e. macrophages) followed by yeast quantification using colony forming unit analysis or flow cytometry. While colony forming unit enumeration has been the most commonly used method in the field, this technique has disadvantages and limitations, including slow growth of some fungal species on solid media and low and/or variable plating efficiencies, which is of particular concern when comparing growth of wild-type and mutant strains. Flow cytometry can provide rapid quantitative information regarding yeast viability, however, adoption of flow cytometric detection for pathogenic yeasts has been limited for a number of practical reasons including its high cost and biosafety considerations. Here, we demonstrate an image-based cytometric methodology using the Cellometer Vision (Nexcelom Bioscience, LLC) for the quantification of viable pathogenic yeasts in co-culture with macrophages. Our studies focus on detection of two human fungal pathogens: Histoplasma capsulatum and Candida albicans. H. capsulatum colonizes alveolar macrophages by replicating within the macrophage phagosome, and here, we quantitatively assess the growth of H. capsulatum yeasts in RAW 264.7 macrophages using acridine orange/propidium iodide staining in combination with image cytometry. Our method faithfully recapitulates growth trends as measured by traditional colony forming unit enumeration, but with significantly increased sensitivity. Additionally, we directly assess infection of live macrophages with a GFP-expressing strain of C. albicans. Our methodology offers a rapid, accurate, and economical means for detection and quantification of important human fungal pathogens in association with host cells.

Here, we demonstrate how image cytometry can be used for quantification of pathogenic fungi in association with host cells in culture. This technique can be used as an alternative to CFU enumeration.

Studies of the cellular pathogenesis mechanisms of pathogenic yeasts such as Candida albicans, Histoplasma capsulatum, and Cryptococcus neoformans ...

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 ...

Electroporation of Functional Bacterial Effectors into Mammalian Cells

The study of protein interactions in the context of living cells can generate critical information about localization, dynamics, and interacting partners. This information is particularly valuable in the context of host-pathogen interactions. Many pathogen proteins function within host cells in a variety of way such as, enabling evasion of the host immune system and survival within the intracellular environment. To study these pathogen-protein host-cell interactions, several approaches are commonly used, including: in vivo infection with a strain expressing a tagged or mutant protein, or introduction of pathogen genes via transfection or transduction. Each of these approaches has advantages and disadvantages. We sought a means to directly introduce exogenous proteins into cells. Electroporation is commonly used to introduce nucleic acids into cells, but has been more rarely applied to proteins although the biophysical basis is exactly the same. A standard electroporator was used to introduce affinity-tagged bacterial effectors into mammalian cells. Human epithelial and mouse macrophage cells were cultured by traditional methods, detached, and placed in 0.4 cm gap electroporation cuvettes with an exogenous bacterial pathogen protein of interest (e.g. Salmonella Typhimurium GtgE). After electroporation (0.3 kV) and a short (4 hr) recovery period, intracellular protein was verified by fluorescently labeling the protein via its affinity tag and examining spatial and temporal distribution by confocal microscopy. The electroporated protein was also shown to be functional inside the cell and capable of correct subcellular trafficking and protein-protein interaction. While the exogenous proteins tended to accumulate on the surface of the cells, the electroporated samples had large increases in intracellular effector concentration relative to incubation alone. The protocol is simple and fast enough to be done in a parallel fashion, allowing for high-throughput characterization of pathogen proteins in host cells including subcellular targeting and function of virulence proteins.

Electroporation was used to insert purified bacterial virulence effector proteins directly into living eukaryotic cells. Protein localization was monitored by confocal immunofluorescence microscopy. This method allows for studies on trafficking, function, and protein-protein interactions using active exogenous proteins, avoiding the need for heterologous expression in eukaryotic cells.

The study of protein interactions in the context of living cells can generate critical information about localization, dynamics, and interacting partners. ...

Porphyromonas gingivalis as a Model Organism for Assessing Interaction of Anaerobic Bacteria with Host Cells

Anaerobic bacteria far outnumber aerobes in many human niches such as the gut, mouth, and vagina. Furthermore, anaerobic infections are common and frequently of indigenous origin. The ability of some anaerobic pathogens to invade human cells gives them adaptive measures to escape innate immunity as well as to modulate host cell behavior. However, ensuring that the anaerobic bacteria are live during experimental investigation of the events may pose challenges. Porphyromonas gingivalis, a Gram-negative anaerobe, is capable of invading a variety of eukaryotic non-phagocytic cells. This article outlines how to successfully culture and assess the ability of P. gingivalis to invade human umbilical vein endothelial cells (HUVECs). Two protocols were developed: one to measure bacteria that can successfully invade and survive within the host, and the other to visualize bacteria interacting with host cells. These techniques necessitate the use of an anaerobic chamber to supply P. gingivalis with an anaerobic environment for optimal growth.
The first protocol is based on the antibiotic protection assay, which is largely used to study the invasion of host cells by bacteria. However, the antibiotic protection assay is limited; only intracellular bacteria that are culturable following antibiotic treatment and host cell lysis are measured. To assess all bacteria interacting with host cells, both live and dead, we developed a protocol that uses fluorescent microscopy to examine host-pathogen interaction. Bacteria are fluorescently labeled with 2&#39;,7&#39;-Bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein acetoxymethyl ester (BCECF-AM) and used to infect eukaryotic cells under anaerobic conditions. Following fixing with paraformaldehyde and permeabilization with 0.2% Triton X-100, host cells are labeled with TRITC phalloidin and DAPI to label the cell cytoskeleton and nucleus, respectively. Multiple images taken at different focal points (Z-stack) are obtained for temporal-spatial visualization of bacteria. Methods used in this study can be applied to any cultivable anaerobe and any eukaryotic cell type.

Porphyromonas gingivalis as a Model Organism for Assessing Interaction of Anaerobic Bacteria with Host Cells

This article presents two protocols: one to measure anaerobic bacteria that can successfully invade and survive within the host, and the other to visualize anaerobic bacteria interacting with host cells. This study can be applied to any cultivable anaerobe and any eukaryotic cell type.

Anaerobic bacteria far outnumber aerobes in many human niches such as the gut, mouth, and vagina. Furthermore, anaerobic infections are common and ...

Analysis of Lymphocyte Extravasation Using an In Vitro Model of the Human Blood-brain Barrier

Lymphocyte extravasation into the central nervous system (CNS) is critical for immune surveillance. Disease-related alterations of lymphocyte extravasation might result in pathophysiological changes in the CNS. Thus, investigation of lymphocyte migration into the CNS is important to understand inflammatory CNS diseases and to develop new therapy approaches. Here we present an in vitro model of the human blood-brain barrier to study lymphocyte extravasation. Human brain microvascular endothelial cells (HBMEC) are confluently grown on a porous polyethylene terephthalate transwell insert to mimic the endothelium of the blood-brain barrier. Barrier function is validated by zonula occludens immunohistochemistry, transendothelial electrical resistance (TEER) measurements as well as analysis of evans blue permeation. This model allows investigation of the diapedesis of rare lymphocyte subsets such as CD56brightCD16dim/- NK cells. Furthermore, the effects of other cells, cytokines and chemokines, disease-related alterations, and distinct treatment regimens on the migratory capacity of lymphocytes can be studied. Finally, the impact of inflammatory stimuli as well as different treatment regimens on the endothelial barrier can be analyzed.

Analysis of Lymphocyte Extravasation Using an In Vitro Model of the Human Blood-brain Barrier

Here, we describe a human blood-brain barrier model enabling to investigate lymphocyte transmigration into the central nervous system in vitro.

Lymphocyte extravasation into the central nervous system (CNS) is critical for immune surveillance. Disease-related alterations of lymphocyte ...

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 ...

High-throughput Measurement of Plasma Membrane Resealing Efficiency in Mammalian Cells

In their physiological environment, mammalian cells are often subjected to mechanical and biochemical stresses that result in plasma membrane damage. In response to these damages, complex molecular machineries rapidly reseal the plasma membrane to restore its barrier function and maintain cell survival. Despite 60 years of research in this field, we still lack a thorough understanding of the cell resealing machinery. With the goal of identifying cellular components that control plasma membrane resealing or drugs that can improve resealing, we have developed a fluorescence-based high-throughput assay that measures the plasma membrane resealing efficiency in mammalian cells cultured in microplates. As a model system for plasma membrane damage, cells are exposed to the bacterial pore-forming toxin listeriolysin O (LLO), which forms large 30-50 nm diameter proteinaceous pores in cholesterol-containing membranes. The use of a temperature-controlled multi-mode microplate reader allows for rapid and sensitive spectrofluorometric measurements in combination with brightfield and fluorescence microscopy imaging of living cells. Kinetic analysis of the fluorescence intensity emitted by a membrane impermeant nucleic acid-binding fluorochrome reflects the extent of membrane wounding and resealing at the cell population level, allowing for the calculation of the cell resealing efficiency. Fluorescence microscopy imaging allows for the enumeration of cells, which constitutively express a fluorescent chimera of the nuclear protein histone 2B, in each well of the microplate to account for potential variations in their number and allows for eventual identification of distinct cell populations. This high-throughput assay is a powerful tool expected to expand our understanding of membrane repair mechanisms via screening for host genes or exogenously added compounds that control plasma membrane resealing.

Here we describe a high-throughput fluorescence-based assay that measures the plasma membrane resealing efficiency through fluorometric and imaging analyses in living cells. This assay can be used for screening drugs or target genes that regulate plasma membrane resealing in mammalian cells.

In their physiological environment, mammalian cells are often subjected to mechanical and biochemical stresses that result in plasma membrane damage. In ...

Image Processing Protocol for the Analysis of the Diffusion and Cluster Size of Membrane Receptors by Fluorescence Microscopy

Particle tracking on a video sequence and the posterior analysis of their trajectories is nowadays a common operation in many biological studies. Using the analysis of cell membrane receptor clusters as a model, we present a detailed protocol for this image analysis task using Fiji (ImageJ) and Matlab routines to: 1) define regions of interest and design masks adapted to these regions; 2) track the particles in fluorescence microscopy videos; 3) analyze the diffusion and intensity characteristics of selected tracks. The quantitative analysis of the diffusion coefficients, types of motion, and cluster size obtained by fluorescence microscopy and image processing provides a valuable tool to objectively determine particle dynamics and the consequences of modifying environmental conditions. In this article we present detailed protocols for the analysis of these features. The method described here not only allows single-molecule tracking detection, but also automates the estimation of lateral diffusion parameters at the cell membrane, classifies the type of trajectory and allows complete analysis thus overcoming the difficulties in quantifying spot size over its entire trajectory at the cell membrane.

Here, we present a protocol for single particle tracking image analysis that allows quantitative evaluation of diffusion coefficients, types of motion and cluster sizes of single particles detected by fluorescence microscopy.

Particle tracking on a video sequence and the posterior analysis of their trajectories is nowadays a common operation in many biological studies. Using ...