The authors thank Ian Clarke and P. Scott Hefty for the pGFP-SW2 and pASK-GFP-mKate2-L2 plasmids, respectively. We thank Paul Miller for technical assistance and Richard Stephens for other resources. This work was funded by NIH grant AI095603 (KH).

1. Generation of Fluorescent Host Cell Lines
<ol>
	<li>Day 1. Plate 293T cells (or other retroviral packaging line) on 6-well plates at 2 x 106 cells/well, to be ~75% confluent the next day. If desired, plate duplicate wells for each retrovirus to be used.</li>
	<li>Day 2. Aspirate cells and add 2 ml fresh growth medium (DMEM + 10% FBS + 2 mM L-glutamine). Transfect cells with 5&#8211;8 &#181;g each of retroviral vector (containing GFP) and packaging vector (e.g., pVSVg) using Lipofectamine 2000 or similar reagent and following the manufacturer&#8217;s instructions.</li>
	<li>Incubate cells with DNA for 4&#8211;6 hr in a 37 &#176;C CO2 incubator, and subsequently replace medium with 1 ml growth medium.</li>
	<li>Incubate cells for 48 hr in 37 &#176;C CO2 incubator.</li>
	<li>Day 3. Plate target cells (e.g., HeLa) on 6-well plates at an approximate density of 105 cells/well, to be ~50% confluent the next day. Verify positive transfection in 293T cells by monitoring GFP fluorescence on an inverted microscope.</li>
	<li>Day 4. Collect retrovirus-containing supernatants off 293T cells using a pipette and filter the supernatant through a 0.45 &#181;m cellulose acetate syringe filter.</li>
	<li>Remove medium from HeLa cells and add 0.5&#160;ml growth medium containing 16 &#181;g/ml polybrene. Add ~0.5 ml of filtered retrovirus to cells drop-wise and incubate cells in a 37 &#176;C CO2 incubator for 24 hr. Store any remaining retrovirus (i.e., from the duplicate well) at 4 &#176;C.</li>
	<li>Day 5. If duplicate wells were made to generate extra retrovirus, repeat step 1.7 to serially infect with the remaining retroviral particles.</li>
	<li>Day 6. Passage transfected HeLa cells 1:1 into a 10 cm dish containing selection antibiotic (e.g., 500 &#181;g/ml neomycin).</li>
	<li>Wait sufficient time for transduced cells to grow back in presence of selection antibiotic, after which cells can be propagated by standard techniques with a lower concentration of selection antibiotic (e.g., 200 &#181;g/ml neomycin). 
	Note: Cells may also be clonally selected for ideal brightness at this time, if necessary.</li>
</ol>
2. Generation of GFP-expressing Chlamydia trachomatis
<ol>
	<li>Prepare 2x CaCl2 buffer (20 mM Tris pH 7.4, 100 mM CaCl2) and 4SP buffer (0.4 M sucrose, 16 mM Na2HPO4).</li>
	<li>Day 1. Thaw frozen C. trachomatis stock on ice and immediately dilute 10 &#181;l bacteria into 50 &#181;l 2x CaCl2 buffer. Add 3&#160;&#181;g plasmid DNA (e.g., pASK-GFP-mKate2-L2), mix well and incubate for 30&#160;min at 25 &#176;C.</li>
	<li>During this step, prepare host cells by trypsinizing a T-75 flask of McCoy cells into a centrifuge tube. Centrifuge the cell suspension at 50 x g for 5 min and discard the supernatant. Resuspend cell pellet in appropriate volume of 1x CaCl2 buffer to give a final concentration of 4 x 107&#160;cells/ml.</li>
	<li>After the 30 min incubation (from step 2.2), add 100 &#181;l prepared McCoy cells to transformation mixture tube (total volume 200 &#181;l) and incubate an additional 20 min at 25 &#176;C. Add 100 &#181;l of this transformed cell mixture to a single well from a 6-well plate and overlay with 2 ml growth medium. Incubate at 37 &#176;C in a CO2 incubator for 2 days.</li>
	<li>Day 3. Lyse McCoy cells infected with C.&#160;trachomatis by replacing medium with 1 ml dH2O and scraping cells with a bent 1,000&#160;&#181;l pipet tip. Add 1 ml 4SP buffer and pass suspension through a 27.5 G needle ~5 times for efficient cell lysis and release of C.&#160;trachomatis.</li>
	<li>Transfer 2 ml of the cell lysate containing C.&#160;trachomatis to a T-75 flask containing a confluent monolayer of freshly plated McCoy cells; add 8 ml DMEM (without FBS) and incubate for 2 hr at 25 &#176;C to allow C.&#160;trachomatis to adhere to cells.</li>
	<li>Aspirate cells and add fresh growth medium supplemented with 10 U/ml penicillin G and 1&#160;&#181;g/ml cycloheximide. Incubate flask for 48 h in a 37 &#176;C CO2 incubator.</li>
	<li>Day 4. Verify by light microscopy that cells are infected and inclusions contain aberrant Chlamydia. Identify inclusions as bright vacuoles on a standard light microscope, and aberrant Chlamydia bodies as ring-like objects within inclusions that are greater than 2 &#181;m diameter.</li>
	<li>Day 5. Lyse culture by replacing medium with 2 ml dH2O and scrape cells into a suspension. Add 2 ml 4SP buffer and pass suspension through a 27.5 G needle ~5 times.</li>
	<li>Use 2 ml of cell lysate to infect a fresh monolayer of McCoy cells in a single well in a 6-well plate. Incubate cells with lysate for 2 hr at 25 &#176;C, aspirate and add fresh growth medium containing penicillin G and cycloheximide.</li>
	<li>Incubate cells in a 37 &#176;C CO2 incubator for 3&#8211;5 days, depending on general health of cells.</li>
	<li>Repeat steps 2.9&#8211;2.10 one or more times, as necessary, passaging cultures into fresh wells in a 6-well plate until normal looking inclusions (devoid of aberrant bodies) have emerged. At this time use an inverted fluorescence microscope to check that C.&#160;trachomatis express mKate2 (red).</li>
	<li>Scale up infected cultures for generation of high titer chlamydial stocks, for example by harvesting Chlamydia transformants from infected McCoy or HeLa cells grown in T-150 flasks.</li>
</ol>
3. Infecting Cells with Chlamydia
<ol>
	<li>Plate GFP-HeLa cells on desired culture vessel, for example glass bottom dishes or chamber slides for high resolution imaging, or 24-well plates for IFU determination and multiwell screening.</li>
	<li>Thaw frozen tube containing C. trachomatis, C. muridarum, or C. pneumoniae on ice and dilute in HBSS to desired multiplicity of infection (MOI), for example MOI = 1.</li>
	<li>Perform either static or centrifugation-aided infections based on the particular Chlamydia strain that will be used.
	<ol>
		<li>For infections with C. trachomatis LGV serovar L2 or C. muridarum, wash GFP-HeLa cells with HBSS and incubate with diluted bacteria for 2 hr at 25 &#176;C. Aspirate and wash cells with HBSS, and incubate cells in growth medium in 37 &#176;C CO2 incubator.</li>
		<li>For infections with C. trachomatis serovar D or C. pneumoniae, wash GFP-HeLa cells with HBSS and add diluted bacteria to cells. Place multiwell plate or glass bottom dish (secured by tape in a multiwell plate lid) in a multiwell plate holder of a swinging bucket rotor attachment in a benchtop centrifuge.</li>
		<li>Centrifuge cells at 900 x g at 25 &#176;C for 1 hr.</li>
		<li>Remove culture vessels from centrifuge and place in a 37 &#176;C CO2 incubator for 1 hr.</li>
		<li>Aspirate cells in a biosafety cabinet, wash cells twice with HBSS, and add fresh growth medium. Place cells in a 37 &#176;C CO2 incubator.</li>
	</ol>
	</li>
</ol>
4.&#160; Live Cell Visualization of Chlamydia Inclusions
<ol>
	<li>Remove Chlamydia infected GFP-HeLa cells from CO2 incubator and replace medium with RPMI without phenol red + 5% FBS.</li>
	<li>Mount cells on an inverted fluorescence microscope (Nikon Eclipse Ti-E or similar) using a 20X or higher oil objective.</li>
	<li>Using imaging software (Volocity 6 or similar), focus on and identify Chlamydia infected GFP-HeLa cells: green cells containing large black holes (Chlamydia inclusions).</li>
	<li>Mark the xyz locations of regions containing Chlamydia-infected cells of interest using imaging software (Volocity 6 or similar). Perform this by manually &#8216;Adding Points&#8217; while in the XY Stage view mode. In this manner, xyz coordinates are saved for each marked location.</li>
	<li>In the Acquisition Setup dialog box, configure software to acquire green (GFP, HeLa cytosol) and red (mKate2, C.&#160;trachomatis) channels, and at a rate of new frames per channel every 5 min.</li>
	<li>Acquire the image sequence using the acquisition protocol entered above by clicking the &#8220;Capture/Record&#8221; button.</li>
</ol>
5. Quantitation of Inclusion Growth Parameters in Live Cells
<ol>
	<li>At desired times post-infection (e.g., 24, 48 hpi) remove Chlamydia infected GFP-HeLa cells from incubator and mount on an inverted fluorescence microscope. Note: A 20X or 40X air objective with a high NA is best-suited for 24-well plates, and a 40X oil objective is recommended for chamber slides. Grow cells on either substrate for this assay.</li>
	<li>Focus on midplanes of cells, where inclusions are at their maximum diameter and yield crisp edges.</li>
	<li>Acquire images for many fields in each well (at least 10 per well); wells typically represent replicates or experimental variables.</li>
	<li>Enumerate the number of inclusions manually, by eye (inclusions are black compartments in GFP-HeLa cells) or use imaging software algorithms to automatically identify and count inclusions.
	<ol>
		<li>Find GFP-HeLa cells by fluorescence intensity (&#8216;Find Objects&#8217; command) and adjust fluorescence threshold intensity based on relative intensities of cells.</li>
		<li>Filter selected objects using size guidelines of keeping those only greater than 5 &#181;m2. This is &#8216;population 1&#8217;.</li>
		<li>Apply &#8216;Fill Holes in Objects&#8217; command using &#8216;population 1&#8217; as the input. This step generates &#8216;population 2&#8217;.</li>
		<li>Subtract &#8216;population 1&#8217; from &#8216;population 2&#8217; to yield positively marked Chlamydia inclusions, designated as &#8216;population 3&#8217;.</li>
		<li>Apply size filter to &#8216;population 3&#8217; (&#8216;Filter Population&#8217; command), for example keeping objects greater than 25 &#181;m2 for inclusions in cells infected for 24 hr or longer. For early infection times, such as prior to 18 hr, set size filter to keep objects greater than 4 &#181;m2, as these inclusions are much smaller. Size filtering results in a population of objects designated as &#8216;inclusions&#8217;.</li>
		<li>Calculate quantitative data from images, for example, inclusion number, inclusion size, and inclusion circumference, using imaging software.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Stamm, W. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chlamydia+trachomatis+infections:+progress+and+problems.">Chlamydia trachomatis infections: progress and problems.</a> The Journal of Infectious Diseases. 179, S380-S383 (1999).</li><li>Schachter, J., Stephens, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Infection+and+disease+epidemiology.">Infection and disease epidemiology.</a> Chlamydia: Intracellular Biology, Pathogenesis, and Immunity. , 139-169 (1999).</li><li>Campbell, L. A., Kuo, C. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chlamydia+pneumoniae--an+infectious+risk+factor+for+atherosclerosis.">Chlamydia pneumoniae--an infectious risk factor for atherosclerosis.</a> Nature Reviews Microbiology. 2 (1), 23-32 (2004).</li><li>Darville, T., Hiltke, T. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pathogenesis+of+genital+tract+disease+due+to+Chlamydia+trachomatis.">Pathogenesis of genital tract disease due to Chlamydia trachomatis.</a> The Journal of Infectious Diseases. 201, S114-S125 (2010).</li><li>Hackstadt, T., Stephens, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+biology.">Cell biology.</a> Chlamydia: Intracellular Biology, Pathogenesis, and Immunity. , 101-138 (1999).</li><li>Wang, Y., Kahane, S., Cutcliffe, L. T., Skilton, R. J., Lambden, P. R., Clarke, I. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+a+transformation+system+for+Chlamydia+trachomatis:+restoration+of+glycogen+biosynthesis+by+acquisition+of+a+plasmid+shuttle+vector).">Development of a transformation system for Chlamydia trachomatis: restoration of glycogen biosynthesis by acquisition of a plasmid shuttle vector).</a> PLoS Pathogens. 7 (9), e1002258 (2011).</li><li>Hackstadt, T., Scidmore, M. A., Rockey, D. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lipid+metabolism+in+Chlamydia+trachomatis-infected+cells:+directed+trafficking+of+Golgi-derived+sphingolipids+to+the+chlamydial+inclusion.">Lipid metabolism in Chlamydia trachomatis-infected cells: directed trafficking of Golgi-derived sphingolipids to the chlamydial inclusion.</a> Proceedings of the National Academy of Sciences of the United States of America. 92 (11), 4877-4881 (1995).</li><li>Boleti, H., Ojcius, D. M., Dautry-Varsat, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+labelling+of+intracellular+bacteria+in+living+host+cells.">Fluorescent labelling of intracellular bacteria in living host cells.</a> Journal of Microbiological Methods. 40 (3), 265-274 (2000).</li><li>Hybiske, K., Stephens, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanisms+of+host+cell+exit+by+the+intracellular+bacterium+Chlamydia.">Mechanisms of host cell exit by the intracellular bacterium Chlamydia.</a> Proceedings of the National Academy of Sciences of the United States of America. 104 (27), 11430-11435 (2007).</li><li>Heinzen, R. A., Hackstadt, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Chlamydia+trachomatis+parasitophorous+vacuolar+membrane+is+not+passively+permeable+to+low-molecular-weight+compounds.">The Chlamydia trachomatis parasitophorous vacuolar membrane is not passively permeable to low-molecular-weight compounds.</a> Infection and Immunity. 65 (3), 1088-1094 (1997).</li><li>Wickstrum, J., Sammons, L. R., Restivo, K. N., Hefty, P. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Conditional+gene+expression+in+Chlamydia+trachomatis+using+the+tet+system.">Conditional gene expression in Chlamydia trachomatis using the tet system.</a> PLoS One. 8 (10), 10-1371 (2013).</li><li>Chin, E., Kirker, K., Zuck, M., James, G., Hybiske, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Actin+recruitment+to+the+Chlamydia+inclusion+is+spatiotemporally+regulated+by+a+mechanism+that+requires+host+and+bacterial+factors.">Actin recruitment to the Chlamydia inclusion is spatiotemporally regulated by a mechanism that requires host and bacterial factors.</a> PLoS One. 7 (10), e46949 (2012).</li><li>Agaisse, H., Derr&#233;, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+C.+trachomatis+cloning+vector+and+the+generation+of+C.+trachomatis+strains+expressing+fluorescent+proteins+under+the+control+of+a+C.+trachomatis+promoter.">A C. trachomatis cloning vector and the generation of C. trachomatis strains expressing fluorescent proteins under the control of a C. trachomatis promoter.</a> PLoS One. 8 (2), e57090 (2013).</li></ol>

The authors declare that they have no competing financial interests.

Here we describe the experimental strategy for generating fluorescent host cells for real time visualization and analysis of Chlamydia inclusions. This vacuole visualization approach confers the powerful ability to illuminate, track and quantitatively measure the dynamic properties of chlamydial inclusions across populations of cells or in single cells over time. Chlamydia inclusions in fluorescent protein labeled cells are strikingly well defined, such that they are easily identified without the need f...

Infectious diseases caused by species of the intracellular bacterium Chlamydia elicit a major burden on global health, including sexually transmitted disease, pelvic inflammatory disease, blindness, pneumonia and possibly atherosclerosis1-4. The ability of Chlamydia to interact with the host cell, from within a vacuole (termed the inclusion), is a critical determinant for their successful infection of cells and the host. The inclusion is a novel pathogenic compartment that enables chlamydial growth and is dynamically modified throughout the entire 2&#8211;3 day developmental cycle of Chlamydia5. The obligate intracellular nature of chlamydiae presents numerous challenges to the research community, in particular for directly studying the unique biology of the inclusion. A major handicap has been the inability to efficiently visualize either intracellular Chlamydia or their inclusion by fluorescent approaches in living cells. A recent discovery has finally revealed the means to generate GFP expressing C. trachomatis6; however, this finding has not yet led to specific labeling of the inclusion. Some techniques have been described for labeling of bacteria and inclusions7,8, but they suffer from shortcomings such as non-specificity, transiency and susceptibility to photobleaching. A key discovery by our group established a new strategy for illuminating the inclusion using GFP expressing host cells9. This strategy rationally exploits the intrinsic impermeability of the inclusion membrane to molecules greater than 520 Da10. When cells are engineered to stably express a particular cytosolic fluorescent protein (e.g., GFP or mCherry), Chlamydia inclusions are visible with remarkable clarity by their complete exclusion of fluorescence. This reverse imaging strategy enables immediate visualization of inclusions for all Chlamydia species and it can be easily adapted for most host cells of interest. As a demonstration of its utility, this method was used previously to reveal and define the cellular exit pathways for Chlamydia spp9.
Here, we further demonstrate how this method is performed, and can be exploited to derive key quantitative data about inclusion growth dynamics. Furthermore, it can effectively substitute for costly antibody-based enumeration methods and can be used in tandem with other fluorescent labels, such as mKate2-expressing Chlamydia11. This powerful combination of tools enables exploration of the physical properties of the chlamydial inclusion membrane inside living host cells.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Lipofectamine 2000</td>
			<td>Invitrogen</td>
			<td>11668</td>
			<td></td>
		</tr>
		<tr>
			<td>Opti-Mem</td>
			<td>Invitrogen</td>
			<td>31985</td>
			<td></td>
		</tr>
		<tr>
			<td>Polybrene</td>
			<td>Sigma</td>
			<td>H9268</td>
			<td></td>
		</tr>
		<tr>
			<td>0.45 &#181;m filters</td>
			<td>Fisher</td>
			<td>09-719D</td>
			<td></td>
		</tr>
		<tr>
			<td>G418</td>
			<td>Invitrogen</td>
			<td>10131</td>
			<td></td>
		</tr>
		<tr>
			<td>HBSS</td>
			<td>Invitrogen</td>
			<td>14025</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>Invitrogen</td>
			<td>11995</td>
			<td></td>
		</tr>
		<tr>
			<td>RPMI 1640</td>
			<td>Invitrogen</td>
			<td>11875</td>
			<td></td>
		</tr>
		<tr>
			<td>RPMI 1640 w/o phenol red</td>
			<td>Cellgro</td>
			<td>17-105-CV</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin G</td>
			<td>Sigma</td>
			<td>13752</td>
			<td></td>
		</tr>
		<tr>
			<td>Cycloheximide</td>
			<td>Sigma</td>
			<td>C7698</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass bottom dishes</td>
			<td>MatTek</td>
			<td>P35G-1.5-14-C</td>
			<td></td>
		</tr>
		<tr>
			<td>Chamber slides, Lab-Tek II</td>
			<td>Nunc</td>
			<td>154526, 154534</td>
			<td></td>
		</tr>
	</tbody>
</table>

using fluorescent proteins to visualize and quantitate chlamydia vacuole growth dynamics in living cells

The obligate intracellular bacterium Chlamydia elicits a great burden on global public health. C.&#160;trachomatis is the leading bacterial cause of sexually transmitted infection and also the primary cause of preventable blindness in the world. An essential determinant for successful infection of host cells by Chlamydia is the bacterium's ability to manipulate host cell signaling from within a novel, vacuolar compartment called the inclusion. From within the inclusion, Chlamydia acquire nutrients required for their 2-3 day developmental growth, and they additionally secrete a panel of effector proteins onto the cytosolic face of the vacuole membrane and into the host cytosol. Gaps in our understanding of Chlamydia biology, however, present significant challenges for visualizing and analyzing this intracellular compartment. Recently, a reverse-imaging strategy for visualizing the inclusion using GFP expressing host cells was described. This approach rationally exploits the intrinsic impermeability of the inclusion membrane to large molecules such as GFP. In this work, we describe how GFP- or mCherry-expressing host cells are generated for subsequent visualization of chlamydial inclusions. Furthermore, this method is shown to effectively substitute for costly antibody-based enumeration methods, can be used in tandem with other fluorescent labels, such as GFP-expressing Chlamydia, and can be exploited to derive key quantitative data about inclusion membrane growth from a range of Chlamydia species and strains.

Mammalian cells expressing cytosolic fluorescent proteins (e.g., GFP) can be engineered to enable illumination of Chlamydia inclusions in live, infected cell cultures. Upon infection with Chlamydia, inclusions are readily visible as black spots in the host cells (Figure 1). The clarity of fluorescence-lacking inclusions can be exploited for the automated identification of inclusions across numerous fields of view and/or treatments (Figure 1). Once fluorescent h...

Watch this Scientific Journal Video about Using Fluorescent Proteins to Visualize and Quantitate Chlamydia Vacuole Growth Dynamics in Living Cells at JoVE.com

Using Fluorescent Proteins to Visualize and Quantitate Chlamydia Vacuole Growth Dynamics in Living Cells

A live cell fluorescent protein based method for illuminating cellular vacuoles (inclusions) containing Chlamydia is described. This strategy enables rapid, automated determination of Chlamydia infectivity in samples and can be used to quantitatively investigate inclusion growth dynamics.

Using Fluorescent Proteins to Visualize and Quantitate Chlamydia Vacuole Growth Dynamics in Living Cells

The obligate intracellular bacterium Chlamydia elicits a great burden on global public health. C.&#160;trachomatis is the leading bacterial cause of ...

using-fluorescent-proteins-to-visualize-quantitate-chlamydia-vacuole

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Immunology and Infection

7.5K visualizações.  University of Washington. A live cell fluorescent protein based method for illuminating cellular vacuoles (inclusions) containing Chlamydia is described. This strategy enables rapid, automated determination of Chlamydia infectivity in samples and can be used to quantitatively investigate inclusion growth dynamics.

Vídeo: Using Fluorescent Proteins to Visualize and Quantitate Chlamydia Vacuole Growth Dynamics in Living Cells

Measuring Calpain Activity in Fixed and Living Cells by Flow Cytometry

Calpains are ubiquitous intracellular, calcium-sensitive, neutral cysteine proteases 1. Calpains play crucial roles in many physiological processes, including signaling, cytoskeletal remodeling, regulation of gene expression, apoptosis and cell cycle progression 1. Calpains have been implicated in many pathologies including muscular dystrophies, cancer, diabetes, Alzheimer's disease and multiple sclerosis 1. Calpain regulation is complex and incompletely understood. mRNA and protein levels correlate poorly with activity, limiting the use of gene or protein expression techniques to measure calpain activity. This video protocol details a flow cytometric assay developed in our laboratory for measuring calpain activity in fixed and living cells. This method uses the fluorescent substrate BOC-LM-CMAC, which is cleaved specifically by calpain, to measure calpain activity. 2 In this video, calpain activity in fixed and living murine 32Dkit leukemia cells, alone or as part of a splenocyte population is measured using an LSRII (BD Bioscience). 32Dkit cells are shown to have elevated activity compared to normal splenocytes.

This article will detail the protocol for measuring calpain activity in fixed and living cells using flow cytometry.

Calpains are ubiquitous intracellular, calcium-sensitive, neutral cysteine proteases 1. Calpains play crucial roles in many physiological processes, ...

Seven Steps to Stellate Cells

Hepatic stellate cells are liver-resident cells of star-like morphology and are located in the space of Disse between liver sinusoidal endothelial cells and hepatocytes1,2. Stellate cells are derived from bone marrow precursors and store up to 80% of the total body vitamin A1, 2. Upon activation, stellate cells differentiate into myofibroblasts to produce extracellular matrix, thus contributing to liver fibrosis3. Based on their ability to contract, myofibroblastic stellate cells can regulate the vascular tone associated with portal hypertension4. Recently, we demonstrated that hepatic stellate cells are potent antigen presenting cells and can activate NKT cells as well as conventional T lymphocytes5. 

 Here we present a method for the efficient preparation of hepatic stellate cells from mouse liver. Due to their perisinusoidal localization, the isolation of hepatic stellate cells is a multi-step process. In order to render stellate cells accessible to isolation from the space of Disse, mouse livers are perfused in situ with the digestive enzymes Pronase E and Collagenase P. Following perfusion, the liver tissue is subjected to additional enzymatic treatment with Pronase E and Collagenase P in vitro. Subsequently, the method takes advantage of the massive amount of vitamin A-storing lipid droplets in hepatic stellate cells. This feature allows the separation of stellate cells from other hepatic cell types by centrifugation on an 8% Nycodenz gradient. The protocol described here yields a highly pure and homogenous population of stellate cells. Purity of preparations can be assessed by staining for the marker molecule glial fibrillary acidic protein (GFAP), prior to analysis by fluorescence microscopy or flow cytometry. Further, light microscopy reveals the unique appearance of star-shaped hepatic stellate cells that harbor high amounts of lipid droplets. 


 Taken together, we present a detailed protocol for the efficient isolation of hepatic stellate cells, including representative images of their morphological appearance and GFAP expression that help to define the stellate cell entity.

Here we describe a method for the isolation of hepatic stellate cells from mouse liver. For stellate cell purification, mouse livers are digested in situ and in vitro by pronase-collagenase treatment prior to density gradient centrifugation. This technique yields highly pure hepatic stellate cells.

Hepatic stellate cells are liver-resident  cells of star-like morphology and are located in the space of Disse between  liver sinusoidal endothelial 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 ...

Measuring Growth and Gene Expression Dynamics of Tumor-Targeted S. Typhimurium Bacteria

The goal of these experiments is to generate quantitative time-course data on the growth and gene expression dynamics of attenuated S. typhimurium bacterial colonies growing inside tumors. 
We generated model xenograft tumors in mice by subcutaneous injection of a human ovarian cancer cell line, OVCAR-8 (NCI DCTD Tumor Repository, Frederick, MD). 
We transformed attenuated strains of S. typhimurium bacteria (ELH430:SL1344 phoPQ- 1) with a constitutively expressed luciferase (luxCDABE) plasmid for visualization2. These strains specifically colonize tumors while remaining essentially non-virulent to the mouse1. 
Once measurable tumors were established, bacteria were injected intravenously via the tail vein with varying dosage. Tumor-localized, bacterial gene expression was monitored in real time over the course of 60 hours using an in vivo imaging system (IVIS). At each time point, tumors were excised, homogenized, and plated to quantitate bacterial colonies for correlation with gene expression data. 
Together, this data yields a quantitative measure of the in vivo growth and gene expression dynamics of bacteria growing inside tumors.

Measuring Growth and Gene Expression Dynamics of Tumor-Targeted S. Typhimurium Bacteria

The goal of these experiments is to generate quantitative time-course data on the growth and gene expression dynamics of attenuated S. typhimurium bacterial colonies growing inside tumors. This video covers tumor cell preparation and implantation, bacteria preparation and injection, whole-animal luminescence imaging, tumor excision, and bacterial colony counting.

The goal of these experiments is to generate quantitative time-course data on the growth and gene expression dynamics of attenuated S. typhimurium ...

Forward Genetic Approaches in Chlamydia trachomatis

Chlamydia trachomatis, the etiological agent of sexually transmitted diseases and ocular infections, remains poorly characterized due to its intractability to experimental transformation with recombinant DNA. We developed an approach to perform genetic analysis in C. trachomatis&#160;despite the lack of molecular genetic tools. Our method involves: i.) chemical mutagenesis to rapidly generate comprehensive libraries of genetically-defined mutants with distinct phenotypes; ii.) whole-genome sequencing (WGS) to map the underlying genetic lesions and to find associations between mutated gene(s) and a common phenotype; iii.) generation of recombinant strains through co-infection of mammalian cells with mutant and wild type bacteria. Accordingly, we were able to establish causal relationships between genotypes and phenotypes. The coupling of chemically-induced gene variation and WGS to establish correlative genotype&#8211;phenotype associations should be broadly applicable to the large list of medically and environmentally important microorganisms currently intractable to genetic analysis.

Forward Genetic Approaches in Chlamydia trachomatis

We describe a methodology to perform genetic analysis in Chlamydia based on chemical mutagenesis and whole genome sequencing. In addition, a system for DNA exchange within infected cells is described that can be used for genetic mapping. This method may be broadly applicable to microbial systems lacking transformation systems and molecular genetic tools.

Chlamydia trachomatis, the etiological agent of sexually transmitted diseases and ocular infections, remains poorly characterized due to its ...

Monitoring Changes in Membrane Polarity, Membrane Integrity, and Intracellular Ion Concentrations in Streptococcus pneumoniae Using Fluorescent Dyes

Membrane depolarization and ion fluxes are events that have been studied extensively in biological systems due to their ability to profoundly impact cellular functions, including energetics and signal transductions. While both fluorescent and electrophysiological methods, including electrode usage and patch-clamping, have been well developed for measuring these events in eukaryotic cells, methodology for measuring similar events in microorganisms have proven more challenging to develop given their small size in combination with the more complex outer surface of bacteria shielding the membrane. During our studies of death-initiation in Streptococcus pneumoniae (pneumococcus), we wanted to elucidate the role of membrane events, including changes in polarity, integrity, and intracellular ion concentrations. Searching the literature, we found that very few studies exist. Other investigators had monitored radioisotope uptake or equilibrium to measure ion fluxes and membrane potential and a limited number of studies, mostly in Gram-negative organisms, had seen some success using carbocyanine or oxonol fluorescent dyes to measure membrane potential, or loading bacteria with cell-permeant acetoxymethyl (AM) ester versions of ion-sensitive fluorescent indicator dyes. We therefore established and optimized protocols for measuring membrane potential, rupture, and ion-transport in the Gram-positive organism S. pneumoniae. We developed protocols using the bis-oxonol dye DiBAC4(3) and the cell-impermeant dye propidium iodide to measure membrane depolarization and rupture, respectively, as well as methods to optimally load the pneumococci with the AM esters of the ratiometric dyes Fura-2, PBFI, and BCECF to detect changes in intracellular concentrations of Ca2+, K+, and H+, respectively, using a fluorescence-detection plate reader. These protocols are the first of their kind for the pneumococcus and the majority of these dyes have not been used in any other bacterial species. Though our protocols have been optimized for S. pneumoniae, we believe these approaches should form an excellent starting-point for similar studies in other bacterial species.

Monitoring Changes in Membrane Polarity, Membrane Integrity, and Intracellular Ion Concentrations in Streptococcus pneumoniae Using Fluorescent Dyes

Unlike that seen for eukaryotes, there is a paucity of studies that detail membrane depolarization and ion concentration changes in bacteria, primarily as their small size makes conventional methods of measurement difficult. Here, we detail protocols for monitoring such events in the significant Gram-positive pathogen Streptococcus pneumoniae utilizing fluorescence techniques.

Membrane depolarization and ion fluxes are events that have been studied extensively in biological systems due to their ability to profoundly impact ...

Using Fluorescent Proteins to Monitor Glycosome Dynamics in the African Trypanosome

Trypanosoma brucei is a kinetoplastid parasite that causes human African trypanosomiasis (HAT), or sleeping sickness, and a wasting disease, nagana, in cattle1. The parasite alternates between the bloodstream of the mammalian host and the tsetse fly vector. The composition of many cellular organelles changes in response to these different extracellular conditions2-5.
Glycosomes are highly specialized peroxisomes in which many of the enzymes involved in glycolysis are compartmentalized. Glycosome composition changes in a developmental and environmentally regulated manner4-11. Currently, the most common techniques used to study glycosome dynamics are electron and fluorescence microscopy; techniques that are expensive, time and labor intensive, and not easily adapted to high throughput analyses.
To overcome these limitations, a fluorescent-glycosome reporter system in which enhanced yellow fluorescent protein (eYFP) is fused to a peroxisome targeting sequence (PTS2), which directs the fusion protein to glycosomes12, has been established. Upon import of the PTS2eYFP fusion protein, glycosomes become fluorescent. Organelle degradation and recycling results in the loss of fluorescence that can be measured by flow cytometry. Large numbers of cells (5,000 cells/sec) can be analyzed in real-time without extensive sample preparation such as fixation and mounting. This method offers a rapid way of detecting changes in organelle composition in response to fluctuating environmental conditions.

Glycosome dynamics in African trypanosomes are difficult to study by traditional cell biology techniques such as electron and fluorescence microscopy. As a means of observing dynamic organelle behavior, a fluorescent-organelle reporter system has been used in conjunction with flow cytometry to monitor real-time glycosome dynamics in live parasites.

Trypanosoma brucei is a kinetoplastid parasite that causes human African trypanosomiasis (HAT), or sleeping sickness, and a wasting disease, nagana, in ...

"Phagosome Closure Assay" to Visualize Phagosome Formation in Three Dimensions Using Total Internal Reflection Fluorescent Microscopy (TIRFM)

Phagocytosis is a mechanism used by specialized cells to internalize and eliminate microorganisms or cellular debris. It relies on profound rearrangements of the actin cytoskeleton that is the driving force for plasma membrane extension around the particle. In addition, efficient engulfment of large material relies on focal exocytosis of intracellular compartments. This process is highly dynamic and numerous molecular players have been described to have a role during phagocytic cup formation. The precise regulation in time and space of all of these molecules, however, remains elusive. In addition, the last step of phagosome closure has been very difficult to observe because inhibition by RNA interference or dominant negative mutants often results in stalled phagocytic cup formation. 
We have set up a dedicated experimental approach using total internal reflection fluorescence microscopy (TIRFM) combined with epifluorescence to monitor step by step the extension of pseudopods and their tips in a phagosome growing around a particle loosely bound to a coverslip. This method allows us to observe, with high resolution the very tips of the pseudopods and their fusion during closure of the phagosome in living cells for two different fluorescently tagged proteins at the same time.

&quot;Phagosome Closure Assay&quot; to Visualize Phagosome Formation in Three Dimensions Using Total Internal Reflection Fluorescent Microscopy TIRFM

We describe an experimental setup to visualize with unprecedented high resolution phagosome formation and closure in three dimensions in living macrophages, using total internal reflection fluorescence microscopy. It allows monitoring of the base of the phagocytic cup, the extending pseudopods, as well as the precise site of phagosome scission.

Phagocytosis is a mechanism used by specialized cells to internalize and eliminate microorganisms or cellular debris. It relies on profound rearrangements ...

Precision-cut Mouse Lung Slices to Visualize Live Pulmonary Dendritic Cells

Inhalation of allergens and pathogens elicits multiple changes in a variety of immune cell types in the lung. Flow cytometry is a powerful technique for quantitative analysis of cell surface proteins on immune cells, but it provides no information on the localization and migration patterns of these cells within the lung. Similarly,&#160;chemotaxis assays can be performed to study the potential of cells to respond to chemotactic factors in vitro, but these assays do not reproduce the complex environment of the intact lung. In contrast to these aforementioned techniques, the location of individual cell types within the lung can be readily visualized by generating Precision-cut Lung Slices (PCLS), staining them with commercially available, fluorescently tagged antibodies, and visualizing the sections by confocal microscopy. PCLS can be used for both live and fixed lung tissue, and the slices can encompass areas as large as a cross section of an entire lobe. We have used this protocol to successfully visualize the location of a wide variety of cell types in the lung, including distinct types of dendritic cells, macrophages, neutrophils, T cells and B cells, as well as structural cells such as lymphatic, endothelial, and epithelial cells. The ability to visualize cellular interactions, such as those between dendritic cells and T cells, in live, three-dimensional lung tissue, can reveal how cells move within the lung and interact with one another at steady state and during inflammation. Thus, when used in combination with other procedures, such as flow cytometry and quantitative PCR, PCLS can contribute to a comprehensive understanding of cellular events that underlie allergic and inflammatory diseases of the lung.

We describe a method for generating Precision-cut Lung Slices (PCLS) and immunostaining them to visualize the localization of various immune cell types in the lung. Our protocol can be extended to visualize the location and function of many different cell types under a variety of conditions.

Inhalation of allergens and pathogens elicits multiple changes in a variety of immune cell types in the lung. Flow cytometry is a powerful technique for ...

Studying Organelle Dynamics in B Cells During Immune Synapse Formation

Recognition of surface-tethered antigens&#160;by the B cell receptor (BCR)&#160;triggers the formation of an immune synapse (IS), where both signaling&#160;and antigen uptake are coordinated. IS formation involves dynamic actin remodeling accompanied by the polarized recruitment to the synaptic membrane of the centrosome and associated intracellular organelles such as lysosomes and the Golgi apparatus. Initial stages of actin remodeling allow B cells to increase their cell surface and maximize the quantity of antigen-BCR complexes gathered at the synapse. Under certain conditions, when B cells recognize antigens associated to rigid surfaces, this process is coupled to the local recruitment and secretion of lysosomes, which can facilitate antigen extraction. Uptaken antigens&#160;are internalized into specialized endo-lysosome compartments for processing into peptides, which are loaded onto major histocompatibility complex II (MHC-II) molecules for further presentation to T helper cells. Therefore, studying organelle dynamics associated with the formation of an IS is crucial to understanding how B cells are activated. In the present article we will discuss both imaging and a biochemical technique used to study changes in intracellular organelle positioning and cytoskeleton rearrangements that are associated with the formation of an IS in B cells.

Herein we describe two approaches to characterize cell polarization events in B lymphocytes during the formation of an IS. The first, involves quantification of organelle recruitment and cytoskeleton rearrangements at the synaptic membrane. The second is a biochemical approach, to characterize changes in composition of the centrosome, which undergoes polarization to the immune synapse.