We would like to acknowledge Mathias S. Dick, Roland F. Dreier and Sebastian R&#252;hl for discussion. This work was supported by an SNSF Professorship PP00P3_139120/1 and a University of Basel project grant ID2153162 to P.B.

1. Digitonin Assay with mCherry-expressing Salmonella (FACS-based Analysis)
<ol>
	<li>Preparing bacteria 
	Note: Salmonella wild-type SL1344 (Streptomycin resistant) expressing mCherry from a plasmid (pFPV encoding mCherry under the rpsM promoter) is used in this assay. Bacterial phagocytosis and proliferation where not altered by the constitutive expression of mCherry (data not shown)&#160;16.
	<ol>
		<li>Inoculate the strain 1 day before the infection in 3 ml of LB medium supplemented with Streptomycin (90 &#181;g/ml) and Ampicillin (100 &#181;g/ml, mCherry expressing vector) in a shaker at 37 &#176;C O/N.</li>
	</ol>
	</li>
	<li>Plating cells for infection.
	<ol>
		<li>Seed wild-type BMDMs in a 24 well plate at a density of 1.25 x 105 cells/well in 1 ml of primary macrophage medium (DMEM supplemented with 10% Fetal Calf Serum (FCS, heat-inactivated, 56 &#176;C, 30 min), 1% HEPES, 1% Non-essential amino acids (NEAA), 1% L-Glutamine and 10% L929 supernatant (conditioned medium from macrophage colony-stimulating factor (M-CSF) producing L929 cells). Seed wells according to Table 1 to account for the individual conditions. Add coverslips to the wells used as controls (Table 1).</li>
		<li>Incubate the cells overnight in an incubator at 37 &#176;C and 5% CO2.</li>
	</ol>
	</li>
</ol>
<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td colspan="2">Permeabilization</td>
			<td colspan="3">Staining</td>
		</tr>
		<tr>
			<td></td>
			<td>Glass coverslip</td>
			<td>Salmonella</td>
			<td>Digitonin</td>
			<td>Saponin</td>
			<td>anti-Salmonella</td>
			<td>anti-Calnexin</td>
			<td>anti-PDI</td>
		</tr>
		<tr>
			<td>A1 (sample)</td>
			<td>-/+*</td>
			<td>+</td>
			<td>+</td>
			<td></td>
			<td>+</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>A2 (sample)</td>
			<td>-/+*</td>
			<td>+</td>
			<td>+</td>
			<td></td>
			<td>+</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>A3 (sample)</td>
			<td>-/+*</td>
			<td>+</td>
			<td>+</td>
			<td></td>
			<td>+</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>A4 (no permeabilization control)</td>
			<td>-/+*</td>
			<td>+</td>
			<td></td>
			<td></td>
			<td>+</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>A5 (complete permeabilization control)</td>
			<td>-/+*</td>
			<td>+</td>
			<td></td>
			<td>+</td>
			<td>+</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>B1 (stained for calnexin)</td>
			<td>+</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td>+</td>
			<td></td>
		</tr>
		<tr>
			<td>B2 (stained for calnexin)</td>
			<td>+</td>
			<td></td>
			<td>+</td>
			<td></td>
			<td></td>
			<td>+</td>
			<td></td>
		</tr>
		<tr>
			<td>B3 (stained for calnexin)</td>
			<td>+</td>
			<td></td>
			<td></td>
			<td>+</td>
			<td></td>
			<td>+</td>
			<td></td>
		</tr>
		<tr>
			<td>C1 (stained for PDI)</td>
			<td>+</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td>+</td>
		</tr>
		<tr>
			<td>C2 (stained for PDI)</td>
			<td>+</td>
			<td></td>
			<td>+</td>
			<td></td>
			<td></td>
			<td></td>
			<td>+</td>
		</tr>
		<tr>
			<td>C3 (stained for PDI)</td>
			<td>+</td>
			<td></td>
			<td></td>
			<td>+</td>
			<td></td>
			<td></td>
			<td>+</td>
		</tr>
	</tbody>
</table>
Table 1: Plating scheme for bone-marrow derived macrophages (BMDMs) in a 24-well format for subsequent infection with Salmonella, permeabilization, and staining. *Depending on whether protocol 1 or protocol 2 is done.
<ol start="3">
	<li>Preparing the bacteria for infection.
	<ol>
		<li>The next day, determine the concentration of Salmonella in the overnight culture by diluting the culture 1:10 in PBS and measuring the OD600 against a PBS only control. This ensures that the OD600 is measured in the linear range of the spectrophotometer.</li>
		<li>Calculate the concentration of Salmonella per ml. An OD600 of 1 corresponds to 1 x 109 bacteria.</li>
		<li>Dilute the bacteria in macrophage medium to reach a Salmonella concentration of 1.25 x 106 cells/ml.</li>
	</ol>
	</li>
	<li>Infection of cells with Salmonella.
	<ol>
		<li>Remove medium from BMDMs. Add 1 ml of macrophage medium to uninfected control wells (B1-B3, C1-C3). Add 1 ml of Salmonella suspension to wells A1 - A5 to reach a multiplicity of infection (m.o.i.) of 10 bacteria per cell.</li>
		<li>Centrifuge the plate for 15 min at 300 x g at 37 &#176;C to synchronize the infection. Transfer the plate in an incubator at 37 &#176;C and 5% CO2.</li>
	</ol>
	</li>
	<li>Killing extracellular Salmonella with Gentamicin.
	<ol>
		<li>At 1 hr post-infection, remove plate from incubator and add 0.1 ml of macrophage medium containing 0.1 mg/ml&#160;Gentamicin to kill extracellular bacteria. Add Gentamicin to all wells, even to uninfected controls, to ensure that all cells are treated the same way. Transfer the plate in an incubator at 37 &#176;C and 5% CO2.</li>
	</ol>
	</li>
	<li>Washing the cells.
	<ol>
		<li>At 2 hr&#160;post infection, remove plate from incubator and wash all wells 2x with 1 ml of plain DMEM (pre-warmed to 37 &#176;C) and replace it with macrophage medium (pre-warmed to 37 &#176;C) containing 10 &#181;g/ml&#160;Gentamycin to prevent growth of any remaining extracellular bacteria. Transfer the plate in an incubator at 37 &#176;C and 5% CO2.</li>
	</ol>
	</li>
	<li>Preparing fresh buffers with detergents and antibodies.
	<ol>
		<li>Just before the desired time point of analysis, prepare a fresh stock solution of digitonin at 1 mg/ml&#160;in KHM Buffer (110 mM potassium acetate, 20 mM HEPES, 2 mM MgCl2, pH 7.3). Filter the stock and dilute in KHM buffer to a working concentration of 50 &#181;g/ml&#160;of digitonin. In addition, prepare a solution of 0.1% Saponin in KHM Buffer, anti-Salmonella antibody coupled to FITC (CSA-1-FITC) at 1/500, anti-calnexin at 1/100 or anti-PDI at 1/100 in KHM Buffer supplemented with 3% BSA.</li>
	</ol>
	</li>
	<li>Cell permeabilization.
	<ol>
		<li>Remove plate from incubator and wash wells 3x&#160;with 0.5 ml of KHM Buffer (pre-warmed to 37 &#176;C). Remove KHM Buffer and add 0.25 ml plain KHM Buffer to wells A4 and B1/C1, KHM Buffer with digitonin to wells A1 - A3 and B2/C2 or KHM Buffer with saponin to wells A5 and B3/C3 (according to Table 1). Incubate for exactly 1 min at RT and immediately wash all wells 3x with 0.5 ml of KHM Buffer (pre-warmed to 37 &#176;C).</li>
	</ol>
	</li>
	<li>Primary antibody staining.
	<ol>
		<li>Remove the KHM Buffer and add the primary antibody solutions (Goat anti Salmonella-FITC, 250 &#181;l/well, 0.1 &#181;g/ml) to the appropriate wells (Figure 1, Table 1). Incubate the plate for 15 min in an incubator at 37 &#176;C and 5% CO2.</li>
		<li>Wash the cells 1x with 1 mL PBS.</li>
	</ol>
	</li>
	<li>Infected Cells (wells A1 - A5): FACS analysis
	<ol>
		<li>Wash cells 3x with PBS and add 0.5 mL of ice-cold PBS containing 0.1% Triton. Transfer to a 5 mL FACS tube with cell strainer cap to break up cell aggregates. Keep samples on ice.</li>
		<li>Analyze the samples of a FACS. Using the fully permeabilized controls, set the gate for total bacteria based on forward and side scatter (FSC/SSC) first, and the mCherry signal (610 nm &#177; 20 nm filter).</li>
		<li>Next, analyze the FITC signal in the total bacterial population using the fully permeabilized and not permeabilized controls to set the gates for cytosolic bacteria (FITC+, mCherry+) and vacuolar bacteria (FITC-, mCherry+) (Figure 3).</li>
		<li>With the gates set, analyze the samples and determine the percentage of cytosolic vs. vacuolar bacteria using FlowJo software according to manufacturer&#8217;s protocol.</li>
	</ol>
	</li>
	<li>Uninfected permeabilization controls (wells B1 - B3, C1 - C3): microscopy
	<ol>
		<li>Fixation
		<ol>
			<li>Remove PBS and add 250 &#181;l of PFA 4% in PBS. Incubate for 10 min at 37 &#176;C.</li>
			<li>Wash wells 2x with 1 ml of PBS and add 250 &#181;l of 0.1 M glycine in PBS for 10 min to quench the fixative. Wash wells 2x with 1 ml of PBS.</li>
		</ol>
		</li>
		<li>Secondary antibody staining.
		<ol>
			<li>Stain permeabilization controls for 1 hr at RT with appropriate secondary antibodies coupled to fluorophores in PBS supplemented with 0.1% Saponin and 3% BSA to fully permeabilize cells.</li>
			<li>Wash coverslips 3x with 1 ml of PBS and mount the coverslips for analysis in mounting medium with DAPI (1.5 &#956;g/ml).</li>
		</ol>
		</li>
		<li>Microscopy analysis.
		<ol>
			<li>Analyze the coverslips with the controls at a 40X or 63X magnification using a confocal fluorescence microscope. If the permeabilization has worked, there should be no staining for non-permeabilized cells, Calnexin staining for both digitonin- and Saponin-permeabilized cells and PDI staining only for saponin-permeabilized cells (Figure 4).</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>



2. Digitonin Assay with Unlabeled Salmonella (Microscopy-based Analysis)



<ol>
	<li>Preparing bacteria for O/N cultures.
	<ol>
		<li>Inoculate unlabeled Salmonella wild-type SL1344 (Streptomycin resistant) 1 day before the infection in 3 ml of LB medium supplemented with Streptomycin (90 &#181;g/ml) in a shaker at 37 &#176;C O/N.</li>
		<li>Plating cells for infection.
		<ol>
			<li>Seed BMDMs in a 24 well plate containing sterile glass coverslips at a density of 1.25 x 105 cells/well in 1 ml of macrophage medium (DMEM supplemented with 10% Fetal Calf Serum (FCS, de-complemented, 56 &#176;C, 30 min), 1% HEPES, 1% L-Non-essential amino acids (NEAA), 1% L-Glutamine and 10% L929 supernatant). Seed wells according to Table 1 to account for the individual conditions.</li>
			<li>Incubate the cells O/N in an incubator at 37 &#176;C and 5% CO2.</li>
		</ol>
		</li>
		<li>Preparing the bacteria for infection.
		<ol>
			<li>The next day, determine the concentration of Salmonella in the O/N culture by diluting the culture 1:10 in PBS and measuring the OD600 against a PBS only control. This ensures that the OD600 is measured in the linear range of the spectrophotometer.</li>
			<li>Determine the concentration of Salmonella per milliliter. An OD600 of 1 corresponds to 1 x 109 bacteria.</li>
			<li>Dilute the bacteria in macrophage medium to reach a Salmonella concentration of 1.25 x 106&#160;cells/ml.</li>
		</ol>
		</li>
		<li>Infection of cells with Salmonella.
		<ol>
			<li>Remove medium from all wells. Add 1 ml of plain macrophage medium to uninfected control wells (B1 - B3, C1 - C3). Add 1 ml of Salmonella suspension to wells A1 - A5 to reach a multiplicity of infection (m.o.i.) of 10 bacteria per cell.</li>
			<li>Centrifuge the plate for 15 min at 300 x g at 37 &#176;C to synchronize the infection. Transfer the plate in an incubator at 37 &#176;C and 5% CO2.</li>
		</ol>
		</li>
		<li>Killing extracellular Salmonella with Gentamycin.
		<ol>
			<li>At 1 hr post-infection, remove plate from incubator and add 0.1 ml of macrophage medium containing 1 mg/ml Gentamycin to kill extracellular bacteria to all wells. Transfer the plate in an incubator at 37 &#176;C and 5% CO2.</li>
		</ol>
		</li>
		<li>Washing the cells.&#160;
		<ol>
			<li>At 2 hr&#160;post-infection, remove plate from incubator and wash each well 3x with 1 ml of fresh macrophage medium containing 10 &#181;g/ml Gentamycin to prevent growth of any remaining extracellular bacteria. Transfer the plate in an incubator at 37&#176;C and 5% CO2.</li>
		</ol>
		</li>
		<li>Preparing fresh buffers with detergents and antibodies.&#160;
		<ol>
			<li>Just before the desired time point of analysis, prepare a fresh stock solution of digitonin at 1 mg/ml in KHM Buffer (110 mM potassium acetate, 20 mM HEPES, 2 mM MgCl2, pH 7.3). Filter the stock and dilute in KHM Buffer to a working concentration of 50 &#181;g/ml of digitonin.</li>
			<li>In addition, prepare a solution of 0.1% Saponin in KHM Buffer, anti-Salmonella-FITC (CSA-1, 0.1 &#181;g/ml), anti-calnexin at 1/100 or anti-PDI at 1/100 in KHM Buffer supplemented with 3% BSA (see materials table for antibody concentration).</li>
		</ol>
		</li>
		<li>Cell permeabilization.
		<ol>
			<li>Remove plate from incubator and wash wells 3x with 0.5 ml of KHM Buffer (pre-warmed to 37 &#176;C). Remove KHM Buffer and add 0.25 ml plain KHM Buffer to wells A4 and B1/C1, KHM Buffer with digitonin to wells A1 - A3 and B2/C2 or KHM Buffer with saponin to wells A5 and B3/C3 (according to Table 1). Incubate for exactly 1 min at RT and immediately wash all wells 3x with 0.5 ml of KHM Buffer (pre-warmed to 37 &#176;C).</li>
		</ol>
		</li>
		<li>Primary antibody staining.
		<ol>
			<li>Remove the KHM Buffer and add the primary antibody solutions (Goat anti Salmonella CSA1-FITC, 250 &#181;l/well, 0.1 &#181;g/ml) to the appropriate wells (Figure 2, Table 1). Incubate the plate for 15 min in an incubator at 37 &#176;C and 5% CO2. Wash 3x with 1 ml PBS.</li>
		</ol>
		</li>
		<li>Fixation.
		<ol>
			<li>Remove PBS and add 250 &#181;l of PFA 4% in PBS. Incubate for 10 min at 37 &#176;C.</li>
			<li>Wash wells 2x with 1 ml of PBS and add 250 &#181;l of 0.1 M Glycine in PBS for 10 min to quench the fixative. Wash wells 2x with 1 ml of PBS.</li>
		</ol>
		</li>
		<li>Total Salmonella staining of infected samples.
		<ol>
			<li>Wash coverslips 3x with PBS with 0.1% Saponin/3% BSA and incubate the samples with an anti-CSA1 antibody (Goat anti-CSA1, 0.1 &#181;g/ml, mouse anti-LPS works as well, 0.1 &#181;g/ml) for 1 hr at RT in PBS with 0.1% Saponin/3% BSA.</li>
			<li>Wash 3x in PBS with 0.1% Saponin/3% BSA. Incubate your samples with a secondary anti-Goat antibody coupled to fluorophore of choice in PBS with 0.1% Saponin/3% BSA for 30 min at RT in the dark.</li>
			<li>Wash coverslips 3x with 1 ml of PBS and mount the coverslips for analysis in mounting medium with DAPI (1.5 &#956;g/ml).&#160;&#160;</li>
		</ol>
		</li>
		<li>Microscopy analysis.
		<ol>
			<li>Acquire data with a confocal microscope by counting cytosolic Salmonella (anti-Salmonella-FITC positive) and total Salmonella (double positive) (Figure 5). Permeabilization controls should show: no staining for non-permeabilized cells, Calnexin staining for digitonin-permeabilized cells and PDI staining for saponin-permeabilized cells (Figure 4). Wells A4 and A5 will serve as internal controls for the Salmonella staining.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

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R., Schlegel, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Membrane+orientation+of+the+human+papillomavirus+type+16+E5+oncoprotein.">Membrane orientation of the human papillomavirus type 16 E5 oncoprotein.</a> J Virol. 84, 1696-1703 (2010).</li><li>Keller, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single+cell+measurements+of+vacuolar+rupture+caused+by+intracellular+pathogens.">Single cell measurements of vacuolar rupture caused by intracellular pathogens.</a> J Vis Exp. , e50116 (2013).</li></ol>

We have nothing to disclose.

Differential permeabilization is an easy and robust method to analyze and quantify the subcellular distribution of bacterial pathogens between vacuolar compartments and the cytosol. The same assay has been used successfully with bacteria such as Francisella novicida2,4 and Shigella flexneri12. However, since many intracellular pathogen alter or modify the host endomembrane structures, the robustness of the digitonin permeabilization will have to be determined on an individual basis...

Intracellular bacterial pathogens replicate either directly in the host cell cytosol, or in specialized vacuolar compartments1. During the initial stage of the infection most pathogens get internalized either by phagocytosis by specialized cells (such as macrophages) or by actively promoting their own uptake into non-phagocytic cells. In phagocytic cells, the phagosome normally fuses with lysosomes to form a degradative compartment, where the phagocytosed particles are digested. Specialized cytosolic bacteria, such as Francisella novicida or Shigella flexneri, escape phagosomal degradation by inducing the rupture of the phagosome and subsequently escape into the cytosol2,3. This requires virulence-associated mechanism such as the Francisella Pathogenicity Island (FPI) or the Shigella flexneri T3SS, which injects effector proteins that promote vacuolar rupture2-4. The host cell cytosol features a number of conserved pattern recognition receptors that normally recognize the presence of pathogens and induce innate immune signaling5. In addition, xenophagy, an anti-microbial form of autophagy can degrade bacteria that enter the cytosol. Cytosolic bacteria are normally well adapted to blunt or circumvent these responses using different strategies. For example, Francisella modifies its LPS to avoid host recognition and Shigella prevents autophagy recruitment using secreted effector proteins6,7.
Another strategy to escape lysosomal degradation is employed by the model vacuolar pathogen Salmonella enterica serovar Typhimurium, thereafter referred to as Salmonella typhimurium. Salmonella uses a T3SS to inject effectors that remodel the initial phagosome into its intracellular niche, the Salmonella-containing vacuole (SCV) 8,9. Continuous manipulation of host pathways is necessary to maintain this vacuole by recruiting lipids and other nutrients to the vacuole. Indeed, a sifA mutant of Salmonella fails to maintain vacuolar stability, and enters the cytosol of host cells within hours after infection, which results in activation of innate immune pathways and autophagy recruitment8. Vacuolar escape of Salmonella varies depending on the cell type that is studies, and several reports have shown that in epithelial cells even WT Salmonella can escape the SCV and hyperreplicate in the cytosol10. Recent reports show that innate immune detection of vacuolar pathogens also depends on the ability of the host to recognize and destabilize pathogen-containing vacuoles (PCVs) 11-14.
Given that both the host or bacteria genotype can affect the distribution of intracellular bacteria between the vacuolar and cytosolic compartment, it is necessary to quantify the number of vacuolar vs. cytosolic bacteria. Since, in most experimental setups host cells are infected with more than one bacterium, and subsequently can harbor several bacteria in different subcellular compartments, a technique is needed that allows quantification on the level of single bacteria. Differential permeabilization (also known as phagosomal protection assay) provides this resolution 2,4,10,12. The assay is based on selective permeabilization of the host cell plasma membrane with the detergent digitonin, which leaves vacuolar membranes intact and thus allows selective staining of cytosolic bacteria with antibodies. Here we provide two protocols for the quantification of cytosolic and vacuolar Salmonella typhimurium using differential permeabilization. The principle of this method is described in Figure 1: Bone-marrow derived macrophages (BMDMs) are infected with stationary phase mCherry-expressing Salmonella. Stationary phase Salmonella need to be used because they downregulate the expression of the SPI-1 T3SS, whose activity would otherwise be recognized by the NLRC4 inflammasome and induce rapid cell death (pyroptosis) of the BMDMs15. Following the infection cells are washed and treated with 50 &#956;g/ml&#160;of digitonin for 1 minute. Cells are washed again immediately and incubated with anti-Salmonella antibodies coupled to FITC to mark bacteria with access to the cytosol. After an additional washing step cells are lysed and the percentage of mCherry+ (vacuolar) and FITC+ / mCherry+ (cytosolic) bacteria is determined by FACS. We also report an adaptation of this method that can be used if no fluorescent protein-expressing strains are available (Figure 2). Additional steps are introduced after the FITC labeling in which cells are fixed and completely permeabilized. Thereafter all bacteria are stained with anti-Salmonella antibodies and corresponding secondary antibodies. Detection is then done by microscopy instead of FACS analysis.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Digitonin</td>
			<td>Sigma</td>
			<td>D5628</td>
			<td>50ug/mL</td>
		</tr>
		<tr>
			<td>PFA</td>
			<td>mpbio</td>
			<td>219998380</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Life Technologies</td>
			<td>15630</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium Acetate</td>
			<td>Sigma</td>
			<td>791733</td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>Sigma</td>
			<td>M8266</td>
			<td></td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Sigma</td>
			<td>A2153</td>
			<td></td>
		</tr>
		<tr>
			<td>anti-Salmonella CSA-1-FITC</td>
			<td>KPL</td>
			<td>01-91-99-MG</td>
			<td>1/500</td>
		</tr>
		<tr>
			<td>anti-Salmonella CSA-1</td>
			<td>KPL</td>
			<td>02-91-99-MG</td>
			<td>1/500</td>
		</tr>
		<tr>
			<td>anti-Calnexin</td>
			<td>Enzo Lifesciences</td>
			<td>SPA-860D</td>
			<td>1/100</td>
		</tr>
		<tr>
			<td>anti-PDI</td>
			<td>Enzo Lifesciences</td>
			<td>SPA-890</td>
			<td>1/100</td>
		</tr>
		<tr>
			<td>Saponin</td>
			<td>Sigma</td>
			<td>47036</td>
			<td></td>
		</tr>
		<tr>
			<td>Vectashield mounting medium</td>
			<td>Vectorlabs</td>
			<td>H-1200</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti Rabbit antibody-488</td>
			<td>Molecular Probes</td>
			<td>A-11070</td>
			<td>1/500</td>
		</tr>
		<tr>
			<td>Glycine</td>
			<td>Sigma</td>
			<td>G8898</td>
			<td></td>
		</tr>
		<tr>
			<td>Gentamicin</td>
			<td>Life Technologies</td>
			<td>15710-49</td>
			<td>100ug/mL and 10ug/mL</td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Promega</td>
			<td>H5141</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>Gibco</td>
			<td>20012-019</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>Sigma</td>
			<td>D6429</td>
			<td></td>
		</tr>
		<tr>
			<td>NEAA</td>
			<td>Amimed</td>
			<td>5-13K00-H</td>
			<td></td>
		</tr>
		<tr>
			<td>LSM700 Confocal microscope</td>
			<td>Zeiss</td>
			<td></td>
			<td>imaging done at 63x</td>
		</tr>
		<tr>
			<td>FACS Fortessa</td>
			<td>BD technologies</td>
			<td></td>
			<td>detection mCherry 610 nm</td>
		</tr>
		<tr>
			<td>FACS Fortessa</td>
			<td>BD technologies</td>
			<td></td>
			<td>detection FITC 530 nm</td>
		</tr>
	</tbody>
</table>

quantification of cytosolic vs. vacuolar salmonella in primary macrophages by differential permeabilization

Intracellular bacterial pathogens can replicate in the cytosol or in specialized pathogen-containing vacuoles (PCVs). To reach the cytosol, bacteria like Shigella flexneri and Francisella novicida need to induce the rupture of the phagosome. In contrast, Salmonella typhimurium replicates in a vacuolar compartment, known as Salmonella-containing vacuole (SCV). However certain mutants of Salmonella fail to maintain SCV integrity and are thus released into the cytosol. The percentage of cytosolic vs. vacuolar bacteria on the level of single bacteria can be measured by differential permeabilization, also known as phagosome-protection assay. The approach makes use of the property of detergent digitonin to selectively bind cholesterol. Since the plasma membrane contains more cholesterol than other cellular membranes, digitonin can be used to selectively permeabilize the plasma membrane while leaving intracellular membranes intact. In brief, following infection with the pathogen expressing a fluorescent marker protein (e.g. mCherry among others), the plasma membrane of host cells is permeabilized with a short incubation in digitonin containing buffer. Cells are then washed and incubated with a primary antibody (coupled to a fluorophore of choice) directed against the bacterium of choice (e.g. anti-Salmonella-FITC) and washed again. If unmarked bacteria are used, an additional step can be done, in which all membranes are permeabilized and all bacteria stained with a corresponding antibody. Following the staining, the percentage of vacuolar and cytosolic bacteria can be quantified by FACS or microscopy by counting single or double-positive events. Here we provide experimental details for use of this technique with the bacterium Salmonella typhimurium. The advantage of this assay is that, in contrast to other assay, it provides a quantification on the level of single bacteria, and if analyzed by microscopy provides the exact number of cytosolic and vacuolar bacteria in a given cell.

Figure 1 and Figure 2 show the schematics of the digitonin assay described in protocol 1 and protocol 2 illustrating the critical steps in the protocol and the results obtained. Figure 3 shows typical FACS results. Positive and negative controls are used to set the gates for mCherry+/FITC- bacteria (vacuolar Salmonella) and mCherry+/FITC+ bacteria (cytosolic Salmonella). Based on these gates the percentage of cytosolic and vacuolar bacteria can be determined in the experime...

Watch this Scientific Journal Video about Quantification of Cytosolic vs. Vacuolar Salmonella in Primary Macrophages by Differential Permeabilization at JoVE.com

Quantification of Cytosolic vs. Vacuolar Salmonella in Primary Macrophages by Differential Permeabilization

We describe the quantification of cytosolic and vacuolar Salmonella typhimurium in bone-marrow derived macrophages using differential digitonin permeabilization.

Quantification of Cytosolic vs. Vacuolar Salmonella in Primary Macrophages by Differential Permeabilization

Intracellular bacterial pathogens can replicate in the cytosol or in specialized pathogen-containing vacuoles (PCVs). To reach the cytosol, bacteria like ...

quantification-cytosolic-vs-vacuolar-salmonella-primary-macrophages

Research

JoVE Journal

Immunology and Infection

10.8K vistas.  University of Basel. We describe the quantification of cytosolic and vacuolar Salmonella typhimurium in bone-marrow derived macrophages using differential digitonin permeabilization.

Video: Quantification of Cytosolic vs. Vacuolar Salmonella in Primary Macrophages by Differential Permeabilization

Measurement of Cytosolic Ca2+ in Isolated Contractile Lymphatics

Lymphatic vessels comprise a multifunctional transport system that maintains fluid homeostasis, delivers lipids to the central circulation, and acts as a surveillance system for potentially harmful antigens, optimizing mucosal immunity and adaptive immune responses1. Lymph is formed from interstitial fluid that enters blind-ended initial lymphatics, and then is transported against a pressure gradient in larger collecting lymphatics. Each collecting lymphatic is made up of a series of segments called lymphangions, separated by bicuspid valves that prevent backflow. Each lymphangion possesses a contractile cycle that propels lymph against a pressure gradient toward the central circulation2. This phasic contractile pattern is analogous to the cardiac cycle, with systolic and diastolic phases, and with a lower contraction frequency4. In addition, lymphatic smooth muscle generates tone and displays myogenic constriction and dilation in response to increases and decreases in luminal pressure, respectively5. A hybrid of molecular mechanisms that support both the phasic and tonic contractility of lymphatics are thus proposed.
Contraction of smooth muscle is generally regulated by the cytosolic Ca2+ concentration ([Ca2+]i) plus sensitivity to Ca2+, of the contractile elements in response to changes in the environment surrounding the cell6. [Ca2+]i is determined by the combination of the movement of Ca2+ through plasma membrane ligand or voltage gated Ca2+ channels and the release and uptake of Ca2+ from internal stores. Cytosolic Ca2+ binds to calmodulin and activates enzymes such as myosin light chain (MLC) kinase (MLCK), which in turn phosphorylates MLC leading to actin-myosin-mediated contraction8. However, the sensitivity of this pathway to Ca2+ can be regulated by the MLC phosphatase (MLCP)9. MLCP activity is regulated by Rho kinase (ROCK) and the myosin phosphatase inhibitor protein CPI-17.
Here, we present a method to evaluate changes in [Ca2+]i over time in isolated, perfused lymphatics in order to study Ca2+-dependent and Ca2+-sensitizing mechanisms of lymphatic smooth muscle contraction. Using isolated rat mesenteric collecting lymphatics we studied stretch-induced changes in [Ca2+]i and contractile activity. The isolated lymphatic model offers the advantage that pressure, flow, and the chemical composition of the bath solution can be tightly controlled. [Ca2+]i was determined by loading lymphatics with the ratiometric, Ca2+-binding dye Fura-2. These studies will provide a new approach to the broader problem of studying the different molecular mechanisms that regulate phasic contractions versus tonic constriction in lymphatic smooth muscle.

Measurement of Cytosolic Ca2+ in Isolated Contractile Lymphatics

We introduce an approach to evaluate the cytosolic Ca2+ concentration in isolated lymphatics to study Ca2+-dependent and Ca2+-sensitizing mechanisms of lymphatic smooth muscle contraction.

Lymphatic vessels comprise a multifunctional transport system that maintains fluid homeostasis, delivers lipids to the central circulation, and acts as a ...

Single Cell Measurements of Vacuolar Rupture Caused by Intracellular Pathogens

Shigella flexneri are pathogenic bacteria that invade host cells entering into an endocytic vacuole. Subsequently, the rupture of this membrane-enclosed compartment allows bacteria to move within the cytosol, proliferate and further invade neighboring cells. Mycobacterium tuberculosis is phagocytosed by immune cells, and has recently been shown to rupture phagosomal membrane in macrophages. We developed a robust assay for tracking phagosomal membrane disruption after host cell entry of Shigella flexneri or Mycobacterium tuberculosis. The approach makes use of CCF4, a FRET reporter sensitive to &beta;-lactamase that equilibrates in the cytosol of host cells. Upon invasion of host cells by bacterial pathogens, the probe remains intact as long as the bacteria reside in membrane-enclosed compartments. After disruption of the vacuole, &beta;-lactamase activity on the surface of the intracellular pathogen cleaves CCF4 instantly leading to a loss of FRET signal and switching its emission spectrum. This robust ratiometric assay yields accurate information about the timing of vacuolar rupture induced by the invading bacteria, and it can be coupled to automated microscopy and image processing by specialized algorithms for the detection of the emission signals of the FRET donor and acceptor. Further, it allows investigating the dynamics of vacuolar disruption elicited by intracellular bacteria in real time in single cells. Finally, it is perfectly suited for high-throughput analysis with a spatio-temporal resolution exceeding previous methods. Here, we provide the experimental details of exemplary protocols for the CCF4 vacuolar rupture assay on HeLa cells and THP-1 macrophages for time-lapse experiments or end points experiments using Shigella flexneri as well as multiple mycobacterial strains such as Mycobacterium marinum, Mycobacterium bovis, and Mycobacterium tuberculosis.

We describe a method for tracking the endomembrane rupture elicited by the intracellular bacteria Shigella flexneri and Mycobacterium tuberculosis upon host cell invasion. Our assay makes use of CCF4, a host cytoplasmic FRET probe in live or fixed cells. This reporter is degraded by an enzyme activity present on the bacterial surface.

Shigella flexneri are pathogenic bacteria that invade host cells entering into an endocytic vacuole. Subsequently, the rupture of this membrane-enclosed ...

High-throughput Assay to Phenotype Salmonella enterica Typhimurium Association, Invasion, and Replication in Macrophages

Salmonella species are zoonotic pathogens and leading causes of food borne illnesses in humans and livestock1. Understanding the mechanisms underlying Salmonella-host interactions are&#160;important to elucidate the molecular pathogenesis of Salmonella infection. The Gentamicin protection assay to phenotype Salmonella association, invasion and replication in phagocytic cells was adapted to allow high-throughput screening to define the roles of deletion mutants of Salmonella enterica serotype Typhimurium in host interactions using RAW 264.7 murine macrophages. Under this protocol, the variance in measurements is significantly reduced compared to the standard protocol, because wild-type and multiple mutant strains can be tested in the same culture dish and at the same time. The use of multichannel pipettes increases the throughput and enhances precision. Furthermore, concerns related to using less host cells per well in 96-well culture dish were addressed. Here, the protocol of the modified in vitro Salmonella invasion assay using phagocytic cells was successfully employed to phenotype 38 individual Salmonella deletion mutants for association, invasion and intracellular replication. The in vitro phenotypes are presented, some of which were subsequently confirmed to have in vivo phenotypes in an animal model. Thus, the modified, standardized assay to phenotype Salmonella association, invasion and replication in macrophages with high-throughput capacity could be utilized more broadly to study bacterial-host interactions.

High-throughput Assay to Phenotype Salmonella enterica Typhimurium Association, Invasion, and Replication in Macrophages

A high-throughput assay to in vitro phenotype Salmonella or other bacterial association, invasion, and replication in phagocytic cells with high-throughput capacity was developed. The method was employed to evaluate Salmonella gene knockout mutant strains for their involvements in host-pathogen interactions.

Salmonella species are zoonotic pathogens and leading causes of food borne illnesses in humans and livestock1. Understanding the mechanisms underlying ...

Highly Efficient Transfection of Human THP-1 Macrophages by Nucleofection

Macrophages, as key players of the innate immune response, are at the focus of research dealing with tissue homeostasis or various pathologies. Transfection with siRNA and plasmid DNA is an efficient tool for studying their function, but transfection of macrophages is not a trivial matter. Although many different approaches for transfection of eukaryotic cells are available, only few allow reliable and efficient transfection of macrophages, but reduced cell vitality and severely altered cell behavior like diminished capability for differentiation or polarization are frequently observed. Therefore a transfection protocol is required that is capable of transferring siRNA and plasmid DNA into macrophages without causing serious side-effects thus allowing the investigation of the effect of the siRNA or plasmid in the context of normal cell behavior. The protocol presented here provides a method for reliably and efficiently transfecting human THP-1 macrophages and monocytes with high cell vitality, high transfection efficiency, and minimal effects on cell behavior. This approach is based on Nucleofection and the protocol has been optimized to maintain maximum capability for cell activation after transfection. The protocol is adequate for adherent cells after detachment as well as cells in suspension, and can be used for small to medium sample numbers. Thus, the method presented is useful for investigating gene regulatory effects during macrophage differentiation and polarization. Apart from presenting results characterizing macrophages transfected according to this protocol in comparison to an alternative chemical method, the impact of cell culture medium selection after transfection on cell behavior is also discussed. The presented data indicate the importance of validating the selection for different experimental settings.

This protocol presents an efficient and reliable method to transfect human THP-1 macrophages with siRNA or plasmid DNA by electroporation with high transfection efficiency while maintaining high cell vitality and full macrophage capacity for differentiation and polarization.

Macrophages, as key players of the innate immune response, are at the focus of research dealing with tissue homeostasis or various pathologies. ...

Proteomic Profiling of Macrophages by 2D Electrophoresis

The goal of the two-dimensional (2D) electrophoresis protocol described here is to show how to analyse the phenotype of human cultured macrophages. The key role of macrophages has been shown in various pathological disorders such as inflammatory, immunological, and infectious diseases. In this protocol, we use primary cultures of human monocyte-derived macrophages that can be differentiated into the M1 (pro-inflammatory) or the M2 (anti-inflammatory) phenotype. This in vitro model is reliable for studying the biological activities of M1 and M2 macrophages and also for a proteomic approach. Proteomic techniques are useful for comparing the phenotype and behaviour of M1 and M2 macrophages during host pathogenicity. 2D gel electrophoresis is a powerful proteomic technique for mapping large numbers of proteins or polypeptides simultaneously. We describe the protocol of 2D electrophoresis using fluorescent dyes, named 2D Differential Gel Electrophoresis (DIGE). The M1 and M2 macrophages proteins are labelled with cyanine dyes before separation by isoelectric focusing, according to their isoelectric point in the first dimension, and their molecular mass, in the second dimension. Separated protein or polypeptidic spots are then used to detect differences in protein or polypeptide expression levels. The proteomic approaches described here allows the investigation of the macrophage protein changes associated with various disorders like host pathogenicity or microbial toxins.

Macrophages are the key cells involved in host pathogenicity. Macrophages display phenotypic and functional diversity that can be analysed and detected by proteomic analysis. This article describes how to perform 2D electrophoresis of primary cultures of human macrophages differentiated into M1 or M2 phenotype.

The goal of the two-dimensional (2D) electrophoresis protocol described here is to show how to analyse the phenotype of human cultured macrophages. The ...

Phagosome Migration and Velocity Measured in Live Primary Human Macrophages Infected with HIV-1

Macrophages are phagocytic cells that play a major role at the crossroads between innate and specific immunity. They can be infected by the human immunodeficiency virus (HIV)-1 and because of their resistance to its cytopathic effects they can be considered to be persistent viral reservoirs. In addition, HIV-infected macrophages exhibit defective functions that contribute to the development of opportunistic diseases.
The exact mechanism by which HIV-1 impairs the phagocytic response of macrophages was unknown. We had previously shown that the uptake of various particulate material by macrophages was inhibited when they were infected with HIV-1. This inhibition was only partial and phagosomes did form within HIV-infected macrophages. Therefore, we focused on analyzing the fate of these phagosomes. Phagosome maturation is accompanied by migration of these compartments towards the cell center, where they fuse with lysosomes, generating phagolysosomes, responsible for degradation of the ingested material. We used IgG-opsonized Sheep Red Blood Cells as a target for phagocytosis. To measure the speed of centripetal movement of phagosomes in individual HIV-infected macrophages, we used a combination of bright field and fluorescence confocal microscopy. We established a method to calculate the distance of phagosomes towards the nucleus, and then to calculate the velocity of the phagosomes. HIV-infected cells were identified thanks to a GFP-expressing virus, but the method is applicable to non-infected cells or any type of infection or treatment.

We describe a method to measure the velocity of phagosomes moving towards the cell center in living cells infected with or without the human immunodeficiency virus (HIV) type 1, using spinning disk confocal fluorescence microscopy to identify fluorescent infected cells and bright field microscopy to detect phagosomes.

Macrophages are phagocytic cells that play a major role at the crossroads between innate and specific immunity. They can be infected by the human ...

Phenotypic Characterization of Macrophages from Rat Kidney by Flow Cytometry

There is increasing evidence suggesting the important role of inflammation and, subsequently, macrophages in the development and progression of renal disease. Macrophages are heterogeneous cells that have been implicated in kidney injury. Macrophages may be classified into two different phenotypes: classically activated macrophages (M1 macrophages), that release pro-inflammatory cytokines and promote fibrosis; and alternatively activated macrophages (M2 macrophages) that are associated with immunoregulatory and tissue-remodeling functions. These macrophage phenotypes need to be discriminated and analyzed to determine their contribution to renal injury. However, there are scarce studies reporting consistent phenotypic and functional information about macrophage subtypes in inflammatory renal disease models, especially in rats. This fact may be related to the limited macrophage markers used in rats, contrary to mice. Therefore, novel strategies are necessary to quantify and characterize the renal content of these infiltrating cells in a reliable way. This manuscript details a protocol for kidney digestion and further phenotypic and quantitative analysis of macrophages from rat kidneys by flow cytometry. Briefly, kidneys were incubated with collagenase and total macrophages were identified according to the dual presence of CD45 (leukocytes common antigen) and CD68 (PAN macrophage marker) in live cells.This was followed by surface staining of CD86 (M1 marker) and CD163 (M2 marker). Rat peritoneal macrophages were used as positive control for macrophage marker detection by flow cytometry. Our protocol resulted in low cellular mortality and allowed characterization of different intracellular and surface protein markers, thus limiting the loss of cellular integrity observed in other protocols. Moreover, this procedure allows the use of macrophages for further techniques, including cell sorting and mRNA or protein expression studies, among others.

This manuscript describes a detailed protocol for phenotypic and quantitative analysis of resident macrophages from rat kidneys by flow cytometry. The resulting stained cells can be also used for other applications, including cell sorting, gene expression analysis or functional studies, thus increasing the information obtained in the experimental model.

There is increasing evidence suggesting the important role of inflammation and, subsequently, macrophages in the development and progression of renal ...

Isolation of Salmonella typhimurium-containing Phagosomes from Macrophages

Salmonella typhimurium is a facultative intracellular bacterium that causes gastroenteritis in humans. After invasion of the lamina propria, S. typhimurium bacteria are quickly detected and phagocytized by macrophages, and contained in vesicles known as phagosomes in order to be degraded. Isolation of S. typhimurium-containing phagosomes have been widely used to study how S. typhimurium infection alters the process of phagosome maturation to prevent bacterial degradation. Classically, the isolation of bacteria-containing phagosomes has been performed by sucrose gradient centrifugation. However, this process is time-consuming, and requires specialized equipment and a certain degree of dexterity. Described here is a simple and quick method for the isolation of S. typhimurium-containing phagosomes from macrophages by coating the bacteria with biotin-streptavidin-conjugated magnetic beads. Phagosomes obtained by this method can be suspended in any buffer of choice, allowing the utilization of isolated phagosomes for a broad range of assays, such as protein, metabolite, and lipid analysis. In summary, this method for the isolation of S. typhimurium-containing phagosomes is specific, efficient, rapid, requires minimum equipment, and is more versatile than the classical method of isolation by sucrose gradient-ultracentrifugation.

Isolation of Salmonella typhimurium-containing Phagosomes from Macrophages

We describe here a simple and quick method for the isolation of Salmonella typhimurium-containing phagosomes from macrophages by coating the bacteria with biotin and streptavidin.

Salmonella typhimurium is a facultative intracellular bacterium that causes gastroenteritis in humans. After invasion of the lamina propria, S. ...

Time-lapse 3D Imaging of Phagocytosis by Mouse Macrophages

Phagocytosis plays a key role in host defense, as well as in tissue development and maintenance, and involves rapid, receptor-mediated rearrangements of the actin cytoskeleton to capture, envelop and engulf large particles. Although phagocytic receptors, downstream signaling pathways, and effectors, such as Rho GTPases, have been identified, the dynamic cytoskeletal remodeling of specific receptor-mediated phagocytic events remain unclear. Four decades ago, two distinct mechanisms of phagocytosis, exemplified by Fc&#947; receptor (Fc&#947;R)- and complement receptor (CR)-mediated phagocytosis, were identified using scanning electron microscopy. Binding of immunoglobulin G (IgG)-opsonized particles to Fc&#947;Rs triggers the protrusion of thin membrane extensions, which initially form a so-called phagocytic cup around the particle before it becomes completely enclosed and retracted into the cell. In contrast, complement opsonized particles appear to sink into the phagocyte following binding to complement receptors. These two modes of phagocytosis, phagocytic cup formation and sinking in, have become well established in the literature. However, the distinctions between the two modes have become blurred by reports that complement receptor-mediated phagocytosis may induce various membrane protrusions. With the availability of high resolution imaging techniques, phagocytosis assays are required that allow real-time 3D (three dimensional) visualization of how specific phagocytic receptors mediate the uptake of individual particles. More commonly used approaches for the study of phagocytosis, such as end-point assays, miss the opportunity to understand what is happening at the interface of particles and phagocytes. Here we describe phagocytic assays, using time-lapse spinning disk confocal microscopy, that allow 3D imaging of single phagocytic events. In addition, we describe assays to unambiguously image Fc&#947; receptor- or complement receptor-mediated phagocytosis.

Here we describe methods using spinning disk confocal microscopy to image single phagocytic events by mouse resident peritoneal macrophages. The protocols can be extended to other phagocytic cells.

Phagocytosis plays a key role in host defense, as well as in tissue development and maintenance, and involves rapid, receptor-mediated rearrangements of ...

Quantification of Antibody-dependent Enhancement of the Zika Virus in Primary Human Cells

The recent emergence of the flavivirus Zika and neurological complications, such as Guillain-Barr&#233; syndrome and microcephaly in infants, has brought serious public safety concerns. Among the risk factors, antibody-dependent enhancement (ADE) poses the most significant threat, as the recent re-emergence of the Zika virus (ZIKV) is primarily in areas where the population has been exposed and is in a state of pre-immunity to other closely related flaviviruses, especially dengue virus (DENV). Here, we describe a protocol for quantifying the effect of human serum antibodies against DENV on ZIKV infection in primary human cells or cell lines.

We describe a method to evaluate the effect of pre-existing immunity against dengue virus on the Zika virus infection by using human serum, primary human cells, and infection quantification by quantitative real-time polymerase chain reaction.

The recent emergence of the flavivirus Zika and neurological complications, such as Guillain-Barr&#233; syndrome and microcephaly in infants, has brought ...