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><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 &amp;micro;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...

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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

Research

JoVE Journal

Immunology and Infection

7.5K Views.  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.

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

Fluorescence Live-cell Imaging of the Complete Vegetative Cell Cycle of the Slow-growing Social Bacterium Myxococcus xanthus

Fluorescence live-cell imaging of bacterial cells is a key method in the analysis of the spatial and temporal dynamics of proteins and chromosomes underlying central cell cycle events. However, imaging of these molecules in slow-growing bacteria represents a challenge due to photobleaching of fluorophores and phototoxicity during image acquisition. Here, we describe a simple protocol to circumvent these limitations in the case of Myxococcus xanthus (which has a generation time of 4 - 6 h). To this end, M. xanthus cells are grown on a thick nutrient-containing agar pad in a temperature-controlled humid environment. Under these conditions, we determine the doubling time of individual cells by following the growth of single cells. Moreover, key cellular processes such as chromosome segregation and cell division can be imaged by fluorescence live-cell imaging of cells containing relevant fluorescently labeled marker proteins such as ParB-YFP, FtsZ-GFP, and mCherry-PomX over multiple cell cycles. Subsequently, the acquired images are processed to generate montages and/or movies.

Fluorescence Live-cell Imaging of the Complete Vegetative Cell Cycle of the Slow-growing Social Bacterium Myxococcus xanthus

Bacterial cells are spatially highly organized. To follow this organization over time in slow growing Myxococcus xanthus cells, a set-up for fluorescence live-cell imaging with high spatiotemporal resolution over several generations was developed. Using this method, spatiotemporal dynamics of important proteins for chromosome segregation and cell division could be determined.

Fluorescence live-cell imaging of bacterial cells is a key method in the analysis of the spatial and temporal dynamics of proteins and chromosomes ...

Biochemistry

Visualizing Intracellular SNARE Trafficking by Fluorescence Lifetime Imaging Microscopy

Soluble N-ethylmaleimide sensitive fusion protein (NSF) attachment protein receptor (SNARE) proteins are key for membrane trafficking, as they catalyze membrane fusion within eukaryotic cells. The SNARE protein family consists of about 36 different members. Specific intracellular transport routes are catalyzed by specific sets of 3 or 4 SNARE proteins that thereby contribute to the specificity and fidelity of membrane trafficking. However, studying the precise function of SNARE proteins is technically challenging, because SNAREs are highly abundant and functionally redundant, with most SNAREs having multiple and overlapping functions. In this protocol, a new method for the visualization of SNARE complex formation in live cells is described. This method is based on expressing SNARE proteins C-terminally fused to fluorescent proteins and measuring their interaction by F&#246;rster resonance energy transfer (FRET) employing fluorescence lifetime imaging microscopy (FLIM). By fitting the fluorescence lifetime histograms with a multicomponent decay model, FRET-FLIM allows (semi-)quantitative estimation of the fraction of the SNARE complex formation at different vesicles. This protocol has been successfully applied to visualize SNARE complex formation at the plasma membrane and at endosomal compartments in mammalian cell lines and primary immune cells, and can be readily extended to study SNARE functions at other organelles in animal, plant, and fungal cells.

This protocol describes a new method allowing for the quantitative visualization of complex formation of SNARE proteins, based on F&#246;rster resonance energy transfer, and fluorescence lifetime imaging microscopy.

Soluble N-ethylmaleimide sensitive fusion protein (NSF) attachment protein receptor (SNARE) proteins are key for membrane trafficking, as they catalyze ...

Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using fluorochromes like Green Fluorescent Protein (GFP) directly attached to the protein of interest has become increasingly popular 1. Using GFP and similar fluorochromes the subcellular localisations and movements of proteins can be detected in a fluorescent microscope. Moreover, also the subnuclear localisation of a certain region of a chromosome can be studied using this technique. GFP is fused to the Lac Repressor protein (LacR) and ectopically expressed in the cell where tandem repeats of the lacO sequence has been inserted into the region of interest on the chromosome2. The LacR-GFP will bind to the lacO repeats and that area of the genome will be visible as a green dot in the fluorescence microscope. Yeast is especially suited for this type of manipulation since homologous recombination is very efficient and thereby enables targeted integration of the lacO repeats and engineered fusion proteins with GFP 3. Here we describe a quantitative method for live cell analysis of fission yeast. Additional protocols for live cell analysis of fission yeast can be found, for example on how to make a movie of the meiotic chromosomal behaviour 4. In this particular experiment we focus on subnuclear organisation and how it is affected during gene induction. We have labelled a gene cluster, named Chr1, by the introduction of lacO binding sites in the vicinity of the genes. The gene cluster is enriched for genes that are induced early during nitrogen starvation of fission yeast 5. In the strain the nuclear membrane (NM) is labelled by the attachment of mCherry to the NM protein Cut11 giving rise to a red fluorescent signal. The Spindle Pole body (SPB) compound Sid4 is fused to Red Fluorescent Protein (Sid4-mRFP) 6. In vegetatively growing yeast cells the centromeres are always attached to the SPB that is embedded in the NM 7. The SPB is identified as a large round structure in the NM. By imaging before and 20 minutes after depletion of the nitrogen source we can determine the distance between the gene cluster (GFP) and the NM/SPB. The mean or median distances before and after nitrogen depletion are compared and we can thus quantify whether or not there is a shift in subcellular localisation of the gene cluster after nitrogen depletion.

The fission yeast, Schizosaccharomyces pombe, is a good model system to study basic cellular processes. Here we describe a method to perform quantitative live cell analysis of fission yeast. In this particular experiment we focus on organisation of the genome within the cell nucleus, but the method can also be used to study cytosolic factors.

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using ...

Biology

Temporal Analysis of the Nuclear-to-cytoplasmic Translocation of a Herpes Simplex Virus 1 Protein by Immunofluorescent Confocal Microscopy

Infected cell protein 0 (ICP0) of herpes simplex virus 1 (HSV-1) is an immediate early protein containing a RING-type E3 ubiquitin ligase. It is responsible for the proteasomal degradation of host restrictive factors and the subsequent viral gene activation. ICP0 contains a canonical nuclear localization sequence (NLS). It enters the nucleus immediately after de novo synthesis and executes its anti-host defense functions mainly in the nucleus. However, later in infection, ICP0 is found solely in the cytoplasm, suggesting the occurrence of a nuclear-to-cytoplasmic translocation during HSV-1 infection. Presumably ICP0 translocation enables ICP0 to modulate its functions according to its subcellular locations at different infection phases. In order to delineate the biological function and regulatory mechanism of ICP0 nuclear-to-cytoplasmic translocation, we modified an immunofluorescent microscopy method to monitor ICP0 trafficking during HSV-1 infection. This protocol involves immunofluorescent staining, confocal microscope imaging, and nuclear vs. cytoplasmic distribution analysis. The goal of this protocol is to adapt the steady state confocal images taken in a time course into a quantitative documentation of ICP0 movement throughout the lytic infection. We propose that this method can be generalized to quantitatively analyze nuclear vs. cytoplasmic localization of other viral or cellular proteins without involving live imaging technology.

ICP0 undergoes nuclear-to-cytoplasmic translocation during HSV-1 infection. The molecular mechanism of this event is not known. Here we describe the use of confocal microscope as a tool to quantify ICP0 movement in HSV-1 infection, which lays the groundwork for quantitatively analyzing ICP0 translocation in future mechanistic studies.

Infected cell protein 0 (ICP0) of herpes simplex virus 1 (HSV-1) is an immediate early protein containing a RING-type E3 ubiquitin ligase. It is ...

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

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

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

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

Monitoring Plasmid Replication in Live Mammalian Cells over Multiple Generations by Fluorescence Microscopy

Few naturally-occurring plasmids are maintained in mammalian cells. Among these are genomes of gamma-herpesviruses, including Epstein-Barr virus (EBV) and Kaposi's Sarcoma-associated herpesvirus (KSHV), which cause multiple human malignancies 1-3. These two genomes are replicated in a licensed manner, each using a single viral protein and cellular replication machinery, and are passed to daughter cells during cell division despite their lacking traditional centromeres 4-8.
Much work has been done to characterize the replications of these plasmid genomes using methods such as Southern blotting and fluorescence in situ hybridization (FISH). These methods are limited, though. Quantitative PCR and Southern blots provide information about the average number of plasmids per cell in a population of cells. FISH is a single-cell assay that reveals both the average number and the distribution of plasmids per cell in the population of cells but is static, allowing no information about the parent or progeny of the examined cell.
Here, we describe a method for visualizing plasmids in live cells. This method is based on the binding of a fluorescently tagged lactose repressor protein to multiple sites in the plasmid of interest 9. The DNA of interest is engineered to include approximately 250 tandem repeats of the lactose operator (LacO) sequence. LacO is specifically bound by the lactose repressor protein (LacI), which can be fused to a fluorescent protein. The fusion protein can either be expressed from the engineered plasmid or introduced by a retroviral vector. In this way, the DNA molecules are fluorescently tagged and therefore become visible via fluorescence microscopy. The fusion protein is blocked from binding the plasmid DNA by culturing cells in the presence of IPTG until the plasmids are ready to be viewed.
This system allows the plasmids to be monitored in living cells through several generations, revealing properties of their synthesis and partitioning to daughter cells. Ideal cells are adherent, easily transfected, and have large nuclei. This technique has been used to determine that 84% of EBV-derived plasmids are synthesized each generation and 88% of the newly synthesized plasmids partition faithfully to daughter cells in HeLa cells. Pairs of these EBV plasmids were seen to be tethered to or associated with sister chromatids after their synthesis in S-phase until they were seen to separate as the sister chromatids separated in Anaphase10. The method is currently being used to study replication of KSHV genomes in HeLa cells and SLK cells. HeLa cells are immortalized human epithelial cells, and SLK cells are immortalized human endothelial cells. Though SLK cells were originally derived from a KSHV lesion, neither the HeLa nor SLK cell line naturally harbors KSHV genomes11. In addition to studying viral replication, this visualization technique can be used to investigate the effects of the addition, removal, or mutation of various DNA sequence elements on synthesis, localization, and partitioning of other recombinant plasmid DNAs.

A method of observing individual DNA molecules in live cells is described. The technique is based on the binding of a fluorescently tagged lac repressor protein to binding sites engineered into the DNA of interest. This method can be adapted to follow many recombinant DNAs in live cells over time.

Few naturally-occurring plasmids are maintained in mammalian cells.  Among these are genomes of gamma-herpesviruses, including Epstein-Barr virus  (EBV) ...

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

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

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

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

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

Visualization of Endoplasmic Reticulum Subdomains in Cultured Cells

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and functions despite the intense interorganelle molecular traffic. The techniques described in this paper are powerful means of studying protein and lipid mobility and trafficking in vivo and in their physiological environment. Fluorescence recovery after photobleaching (FRAP) and fluorescence loss in photobleaching (FLIP) are widely used live-cell imaging techniques for studying intracellular trafficking through the exo-endocytic pathway, the continuity between organelles or subcompartments, the formation of protein complexes, and protein localization in lipid microdomains, all of which can be observed under physiological and pathological conditions. The limitations of these approaches are mainly due to the use of fluorescent fusion proteins, and their potential drawbacks include artifactual over-expression in cells and the possibility of differences in the folding and localization of tagged and native proteins. Finally, as the limit of resolution of optical microscopy (about 200 nm) does not allow investigation of the fine structure of the ER or the specific subcompartments that can originate in cells under stress (i.e. hypoxia, drug administration, the over-expression of transmembrane ER resident proteins) or under pathological conditions, we combine live-cell imaging of cultured transfected cells with ultrastructural analyses based on transmission electron microscopy.

We describe the imaging approaches we use to investigate the distribution and mobility of the transfected fluorescent proteins resident in the endoplasmic reticulum (ER) by means of the confocal imaging of living cells. We also ultrastructurally analyze the effect of their expression on the architecture of this subcellular compartment.

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and ...

Applications of pHluorin for Quantitative, Kinetic and High-throughput Analysis of Endocytosis in Budding Yeast

Green fluorescent protein (GFP) and its variants are widely used tools for studying protein localization and dynamics of events such as cytoskeletal remodeling and vesicular trafficking in living cells. Quantitative methodologies using chimeric GFP fusions have been developed for many applications; however, GFP is somewhat resistant to proteolysis, thus its fluorescence persists in the lysosome/vacuole, which can impede quantification of cargo trafficking in the endocytic pathway. An alternative method for quantifying endocytosis and post-endocytic trafficking events makes use of superecliptic pHluorin, a pH-sensitive variant of GFP that is quenched in acidic environments. Chimeric fusion of pHluorin to the cytoplasmic tail of transmembrane cargo proteins results in a dampening of fluorescence upon incorporation of the cargo into multivesicular bodies (MVBs) and delivery to the lysosome/vacuole lumen. Thus, quenching of vacuolar fluorescence facilitates quantification of endocytosis and early events in the endocytic pathway. This paper describes methods using pHluorin-tagged cargos for quantification of endocytosis via fluorescence microscopy, as well as population-based assays using flow cytometry.

Accurate quantification of vesicular trafficking events often provides key insights into roles for specific proteins and the effects of mutations. This paper presents methods for using superecliptic pHluorin, a pH-sensitive GFP variant, as a tool for quantification of endocytic events in living cells using quantitative fluorescence microscopy and flow cytometry.

Green fluorescent protein (GFP) and its variants are widely used tools for studying protein localization and dynamics of events such as cytoskeletal ...

Live Cell Fluorescence Microscopy to Observe Essential Processes During Microbial Cell Growth

Core cellular processes such as DNA replication and segregation, protein synthesis, cell wall biosynthesis, and cell division rely on the function of proteins which are essential for bacterial survival. A series of target-specific dyes can be used as probes to better understand these processes. Staining with lipophilic dyes enables the observation of membrane structure, visualization of lipid microdomains, and detection of membrane blebs. Use of fluorescent-d-amino acids (FDAAs) to probe the sites of peptidoglycan biosynthesis can indicate potential defects in cell wall biogenesis or cell growth patterning. Finally, nucleic acid stains can indicate possible defects in DNA replication or chromosome segregation. Cyanine DNA stains label living cells and are suitable for time-lapse microscopy enabling real-time observations of nucleoid morphology during cell growth. Protocols for cell labeling can be applied to protein depletion mutants to identify defects in membrane structure, cell wall biogenesis, or chromosome segregation. Furthermore, time-lapse microscopy can be used to monitor morphological changes as an essential protein is removed and can provide additional insights into protein function. For example, the depletion of essential cell division proteins results in filamentation or branching, whereas the depletion of cell growth proteins may cause cells to become shorter or rounder. Here, protocols for cell growth, target-specific labeling, and time-lapse microscopy are provided for the bacterial plant pathogen Agrobacterium tumefaciens. Together, target-specific dyes and time-lapse microscopy enable characterization of essential processes in A. tumefaciens. Finally, the protocols provided can be readily modified to probe essential processes in other bacteria.

Understanding the function of essential processes in bacteria is challenging. Fluorescence microscopy with target-specific dyes can provide key insights into microbial cell growth and cell cycle progression. Here, Agrobacterium tumefaciens is used as a model bacterium to highlight methods for live cell imaging for characterization of essential processes.

Core cellular processes such as DNA replication and segregation, protein synthesis, cell wall biosynthesis, and cell division rely on the function of ...

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

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