Name: kinetic visualization of single-cell interspecies bacterial interactions
Uploaded: 2020-08-05T13:30:00
Description: Watch this Scientific Journal Video about Kinetic Visualization of Single-Cell Interspecies Bacterial Interactions at JoVE.com

This work was supported by funding from the Cystic Fibrosis Foundation Postdoc-to-Faculty Transition Award LIMOLI18F5 (DHL), Cystic Fibrosis Foundation Junior Faculty Recruitment Award LIMOLI19R3 (DHL), and NIH T32 Training Grant 5T32HL007638-34 (ASP). We thank Jeffrey Meisner, Minsu Kim, and Ethan Garner for sharing initial protocols and advice for imaging and making pads.

NOTE: A full description and catalog numbers for all supplies in this protocol can be found in the Table of Materials.
1. Preparation of M8T minimal media
<ol>
	<li>Prepare 1 L of M8 minimal salts base (5x) by dissolving 64 g of Na2HPO4&#183;7H2O, 15 g of KH2PO4, and 2.5 g NaCl in 800 mL of ultrapure water (UPW, resistivity 18 M&#937;/cm) in a 2 L bottle. pH to 7.6. Complete volume to 1 L with UPW. Autoclave for 45 min.</li>
	<li>Prepare...

<ol>
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R., Lawrence, J. R., Sutton, B., Caldwell, D. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+laminar+flow+velocity+on+the+kinetics+of+surface+recolonization+by+Mot++and+Mot-+Pseudomonas+fluorescens.">Effect of laminar flow velocity on the kinetics of surface recolonization by Mot+ and Mot- Pseudomonas fluorescens.</a> Microbial Ecology. 18, 1-19 (1989).</li><li>Lawrence, J. R., Korber, D. R., Caldwell, D. 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The methods presented here describe a protocol for live-cell imaging of bacterial species interactions at the single-cell level with modifications for other applications including cell tracking and monitoring cell viability. This method opens new avenues for studying single-cell behaviors of microorganisms in coculture with other species over time. Specifically, the protocol demonstrates the usefulness of this coculture method in observing bacterial surface behaviors, particularly when studying organisms that have both s...

Polymicrobial communities are common&#160;in nature, spanning a variety of environmental1,2,3 and human niches4,5. Some of the most notorious polymicrobial infections in human disease include dental biofilms6, chronic wounds7,8, and respiratory infections in individuals with chronic obstructive pulmonary disease9, ventilator-associated pneumonia10, and the genetic disease cystic fibrosis (CF)11,12. These infections are often composed of diverse microbial species; however, a recent focus on the interactions between the Gram-positive bacterium Staphylococcus aureus and the Gram-negative bacterium Pseudomonas aeruginosa has emerged. Studies including patients coinfected with these organisms and in vitro analyses reveal both competitive and cooperative interactions that can have profound and unexpected influences on disease progression, microbial survival, virulence, metabolism, and antibiotic susceptibility (reviewed in detail by Hotterbeekx et al. 201713 and Limoli and Hoffman 20193).
The growing interest in interspecies interactions during infection has yielded a variety of methods for studying polymicrobial community behaviors. Typically, these studies have utilized plate or broth-based assays to investigate the phenotypic differences between monoculture and coculture. For example, coculturing P. aeruginosa and S. aureus on solid surfaces has allowed for visualization of growth inhibition or changes in colony phenotype, pigment, or polysaccharide production14,15,16. Mixed species biofilms, on biotic or abiotic surfaces, as well as coculturing bacterial species in liquid culture also has enabled measurement of changes in growth, metabolism, antibiotic tolerance, competition and viability, in addition to gene and protein expression17,18. While these bulk culture experiments have revealed insight into how P. aeruginosa and S. aureus might influence one another on the community-scale, they are unable to answer important questions about the interactions that take place at the single-cell level. The method presented here adds to the approaches for studying interspecies interactions by focusing on the changes in movement, cell viability, and gene expression of single cells within a cocultured community over time.
Single-cell interactions can widely differ from the interactions that take place in a bulk community of cells. Through single-cell analysis, heterogeneity within a community can be quantified to study how spatial placement of cells influence community dynamics or how gene and protein expression levels change within individual members of a group. Tracking cells can also provide insight into how single cells move and behave over time, through multiple generations. By tracking cell movement and changes in gene expression concurrently, correlations can be made between gene fluctuations and corresponding phenotypes. Previous protocols for studying individual bacterial species at the single-cell level have been described. These studies take advantage of live-imaging cells over time in a single plane, and have been useful for observing phenotypes like cell division and antibiotic susceptibility19,20. Additional live-imaging microscopy has been utilized to monitor growth, motility, surface colonization and biofilm formation of single bacterial species21,22,23. However, while these studies have been insightful for understanding the physiology of bacteria in monoculture, experiments for observing single-cell behavior of multiple bacterial species over time in coculture are limited.
Here, previous protocols used for single-species imaging have been adapted to study interactions between P. aeruginosa and S. aureus. These organisms can be differentiated under phase contrast based on their cell morphologies (P. aeruginosa are bacilli&#160;and S. aureus are cocci). Development of this method recently enabled the visualization of previously undescribed motility behaviors of P. aeruginosa in the presence of S. aureus24. P. aeruginosa was found to be capable of sensing S. aureus from a distance and responding with increased and directional single-cell movements towards clusters of S. aureus cells. P. aeruginosa movement towards S. aureus was found to require the type IV pili (TFP), hair-like projections whose coordinated extension and retraction generate a movement called twitching motility25.
These studies demonstrate the utility of this method for interrogating earlier interactions between species. However, imaging at high cell densities at later interaction time points is difficult given that single layers of cells can no longer be identified, which mostly poses issues during post-imaging analysis. Given this limitation, the method is best suited for earlier interactions that could then be followed up with traditional macroscopic assays at higher cell densities representative of later interactions. Additional limitations of this method include the low-throughput nature, since only one sample can be imaged at a time and the cost of the microscope, camera, and environmental chamber. Moreover, fluorescence microscopy poses risks to the bacterial cells like phototoxicity and photobleaching, therefore limiting the frequency that fluorescence images can be acquired. Lastly, the agarose pads used in this method are highly susceptible to changes in the environment, making it critical to control for conditions like temperature and humidity, given that the pads can start to shrink or expand if the conditions are not correct. Finally, while this method does not mimic the host environment, it provides clues for how different bacterial species respond on surfaces, which can be followed-up in assays designed to mimic environmental/host conditions.
This method differs from previous studies tracking single-cell movement, in that cells are inoculated between a coverslip and agarose pad, restricting cells to the surface. This enables cell tracking over time in a single plane; however, limits cycles of transient surface engagement observed when cells are submerged in liquid26. An additional benefit of imaging bacteria under an agarose pad is that it mimics macroscopic plate-based sub-surface inoculation assays classically used to examine P. aeruginosa twitching motility25. In this assay, bacterial cells are inoculated between the bottom of a Petri dish and the agar, keeping the cells in a single plane as they move across the bottom of the dish outward from the point of inoculation, much like this microscopy protocol.
The time-lapse microscopy protocol for visualizing interspecies interactions presented here consists of 1) preparing the bacterial sample and agarose pad, 2) selecting microscope settings for imaging acquisition and 3) post-imaging analysis. Detailed visualization of cell movement and tracking can be performed by acquisition of images at short time intervals by phase contrast. Fluorescence microscopy can also be utilized to determine cell viability or gene expression over time. Here, we show one example of adaptation for fluorescence microscopy through the addition of viability dyes to the agarose pads.

<table>
	<tbody>
		<tr>
			<td>Agarose pads</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>35 mm Glass Bottom Dish with 20 mm Micro-well #1.5 Cover Glass</td>
			<td>Cellvis</td>
			<td>D35-20-1.5-N</td>
			<td>One for agarose pad molds, one for experiment</td>
		</tr>
		<tr>
			<td>KimWipes</td>
			<td>Kimberly-Clark Professional</td>
			<td>06-666A</td>
			<td></td>
		</tr>
		<tr>
			<td>Low-Melt Agarose</td>
			<td>Nu-Sieve GTG/Lonza</td>
			<td>50081</td>
			<td>For making agarose pads</td>
		</tr>
		<tr>
			<td>Round-Bottom Spatulas</td>
			<td>VWR</td>
			<td>82027-492</td>
			<td></td>
		</tr>
		<tr>
			<td>Round-Tapered Spatulas</td>
			<td>VWR</td>
			<td>82027-530</td>
			<td></td>
		</tr>
		<tr>
			<td>Silicon Isolators, Press-to-Seal, 1 well, D diameter 2.0 mm 20 mm, silicone/adhesive</td>
			<td>Sigma-Aldrich</td>
			<td>S6685-25EA</td>
			<td>For agarose pad molds</td>
		</tr>
		<tr>
			<td>Sterile Petri Plates, 85 mm</td>
			<td>Kord-Valmark /sold by RPI</td>
			<td>2900</td>
			<td></td>
		</tr>
		<tr>
			<td>Tweezers</td>
			<td>VWR</td>
			<td>89259-944</td>
			<td></td>
		</tr>
		<tr>
			<td>M8T Minimal Media</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>D (+) Glucose</td>
			<td>RPI</td>
			<td>G32045</td>
			<td></td>
		</tr>
		<tr>
			<td>KH2PO4</td>
			<td>RPI</td>
			<td>P250500</td>
			<td></td>
		</tr>
		<tr>
			<td>MgSO4</td>
			<td>Sigma-Aldrich</td>
			<td>208094</td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>RPI</td>
			<td>S23025</td>
			<td></td>
		</tr>
		<tr>
			<td>Na2HPO4.7H2O</td>
			<td>Sigma-Aldrich</td>
			<td>230391</td>
			<td></td>
		</tr>
		<tr>
			<td>Tryptone</td>
			<td>BD Biosciences</td>
			<td>DF0123173</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Andor Sona 4.2B-11</td>
			<td>Andor</td>
			<td>77026135</td>
			<td>Camera. 4.2 Megapixel Back-illuminated sCMOS, 11 &#956;m pixel, 95% QE, 48 fps, USB 3.0, F-mount.</td>
		</tr>
		<tr>
			<td>Filter Cube GFP</td>
			<td>Nikon</td>
			<td>96372</td>
			<td>Filter cube</td>
		</tr>
		<tr>
			<td>Filter Cube TxRed</td>
			<td>Nikon</td>
			<td>96375</td>
			<td>Filter cube</td>
		</tr>
		<tr>
			<td>H201-NIKON-TI-S-ER</td>
			<td>Okolab</td>
			<td>77057447</td>
			<td>Stagetop incubator</td>
		</tr>
		<tr>
			<td>Nikon NIS-Elements AR with GA3 and 2D and 3D tracking</td>
			<td>Nikon</td>
			<td>77010609, MQS43110, 77010603, MQS42950</td>
			<td>Software for data analysis</td>
		</tr>
		<tr>
			<td>Nikon Ti2 Eclipse</td>
			<td>Nikon</td>
			<td>Model Ti2-E</td>
			<td>Microscope</td>
		</tr>
		<tr>
			<td>CFI Plan Apo &#411;20x objective (0.75NA)</td>
			<td>Nikon</td>
			<td>MRD00205</td>
			<td>Objective</td>
		</tr>
		<tr>
			<td>CFI Plan Apo &#411;100x oil Ph3 DM objective (1.45NA)</td>
			<td>Nikon</td>
			<td>MRD31905</td>
			<td>Objective</td>
		</tr>
		<tr>
			<td>ThermoBox with built-in fan heaters</td>
			<td>Tokai Hit</td>
			<td>TI2TB-E-BK</td>
			<td>Enclosure</td>
		</tr>
		<tr>
			<td>Bacterial Strains</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Pseudomonas aeruginosa PA14 (WT)</td>
			<td></td>
			<td>PMID: 7604262</td>
			<td>Non-mucoid prototroph</td>
		</tr>
		<tr>
			<td>Pseudomonas aeruginosa PA14 (WT) pSMC21 (Ptac-GFP)</td>
			<td></td>
			<td>PMID: 9361441</td>
			<td></td>
		</tr>
		<tr>
			<td>Pseudomonas aeruginosa PAO1 (WT) pPrpoD-mKate2</td>
			<td></td>
			<td>PMID: 26041805</td>
			<td></td>
		</tr>
		<tr>
			<td>Staphylococcus aureus USA300 LAC (WT)</td>
			<td></td>
			<td>PMID: 23404398</td>
			<td>USA300 CA-Methicillin resistant strain LAC without plasmids</td>
		</tr>
		<tr>
			<td>Staphylococcus aureus USA300 LAC (WT) pCM29 (sarAP1-sGFP)</td>
			<td></td>
			<td>PMID: 20829608</td>
			<td></td>
		</tr>
		<tr>
			<td>Staphylococcus aureus USA300 LAC &#9651;agrBDCA</td>
			<td></td>
			<td>PMID: 31713513</td>
			<td></td>
		</tr>
		<tr>
			<td>Viability Stain</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Propidium Iodide</td>
			<td>Invitrogen</td>
			<td>L7012</td>
			<td>LIVE/DEAD&#8482; BacLight&#8482; Bacterial Viability Kit</td>
		</tr>
	</tbody>
</table>

kinetic visualization of single-cell interspecies bacterial interactions

Polymicrobial communities are ubiquitous in nature, yet studying their interactions at the single-cell level is difficult. Thus, a microscopy-based method has been developed for observing interspecies interactions between two bacterial pathogens. The use of this method to interrogate interactions between a motile Gram-negative pathogen, Pseudomonas aeruginosa and a non-motile Gram-positive pathogen, Staphylococcus aureus is demonstrated here. This protocol consists of co-inoculating each species between a coverslip and an agarose pad, which maintains the cells in a single plane and allows for visualization of bacterial behaviors in both space and time.
Furthermore, the time-lapse microscopy demonstrated here is ideal for visualizing the early interactions that take place between two or more bacterial species, including changes in bacterial species motility in monoculture and in coculture with other species. Due to the nature of the limited sample space in the microscopy setup, this protocol is less applicable for studying later interactions between bacterial species once cell populations are too high. However, there are several different applications of the protocol which include the use of staining for imaging live and dead bacterial cells, quantification of gene or protein expression through fluorescent reporters, and tracking bacterial cell movement in both single species and multispecies experiments.

Successful use of the described method will result in a series of frames that generate a video in which the interspecies interactions can be observed over time. The schematic in Figure 1 provides a visual to highlight the key steps involved in preparing materials for imaging. Use of this method has allowed the demonstration of P. aeruginosa cells exhibiting different behaviors in monoculture versus in coculture with S. aureus. Compared to the P. aeruginosa cells in...

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Kinetic Visualization of Single-Cell Interspecies Bacterial Interactions

This live-bacterial cell imaging protocol allows for visualization of interactions between multiple bacterial species at the single-cell level over time. Time-lapse imaging allows for observation of each bacterial species in monoculture or coculture to interrogate interspecies interactions in multispecies bacterial communities, including individual cell motility and viability.

Polymicrobial communities are ubiquitous in nature, yet studying their interactions at the single-cell level is difficult. Thus, a microscopy-based method ...

This protocol allows for visualization and tracking of interspecies bacterial interactions at the single cell level over time. This techniques main advantage is its applicability to studying bacterial interactions, including cell tracking, gene expression and viability staining. Additionally, the conditions can be modified to study different organisms.To begin, prepare agarose pads by melting 2%low melt agarose in 10 milliliters of bacterial growth media in a clean 50 milliliter Erlenmeyer flask. Microwave the flask in short intervals until the agarose is in solution, then let it cool in a 50 degree Celsius water bath for at least 15 minutes. Align a silicon cut out with the opening in a 35 millimeter dish, then flip the dish and lightly tap the silicon cut out with a spatula to secure it to the dish and to remove any air bubbles.Once the agarose has cooled, pipette 915 microliters of molten agarose into the mold and let the pad dry. Warm the microscope environmental chamber to the experimental temperature at least two hours prior to setting up the experiment. Prepare humidity wipes by tightly rolling up a lint-free wipe.Place the rolled wipe inside a sterile Petri dish and add 500 microliters of sterile water directly to the wipes. Warm the wipes, pads and a sterile 35 millimeter dish to the experimental temperature in order to equilibrate all materials. Once the pads are dry, pipette one microliter of co-culture evenly across the bottom of a pre-warmed sterile 35 millimeter glass coverslip dish.Remove the silicone cut out from the agarose mold using sterile tweezers, tilt it to the side and slip a slightly bent spatula under the edge of the agarose pad to drop it onto a sterile petriplate lid. Transfer the pad to the dish with the bacterial cells bottom side down by sliding a 90 degree angled spatula under the pad and placing it on top of the inoculated cover slip. Use the spatula to make the pad flush against the cover slip and gently press out any air bubbles.Remove excess moisture from the moist wipe and place it around the edge of the dish, making sure it does not touch the pad. The sample is now ready for imaging. Prior to inoculated pads, set up the microscope by performing color illumination to align all image frames.Once the inoculated dish is ready, view the dish with 100X objective and focus on the bacterial results. Click the phase option in the imaging software and adjust the percentage of DIA LED light emitted by selecting the light source and sliding the bar on the light percentage scale. Alternatively, manually enter an exposure time.Save the settings for the phase channel by right clicking on the phase channel and selecting the Assign Current Microscope Setting option. If using fluorescence, select the fluorescence channel of interest and set the percentage of fluorescence light emitted, then adjust the exposure time as previously described. Alternatively, change the bid depth to adjust the dynamic range by selecting one of the other options in the drop down menu.Save the settings for the fluorescence channel by right clicking on the fluorescence channel and selecting the Assign Current Microscope Setting option. In the XY tab, click the box in the point name column to save the selected XY position of interest. Then click the PFS box at the bottom of the tab to turn on the perfect focus system.Focus the cells and click the arrow in the PFS column to save the focus of the XY position. Repeat this process for all other XY positions. Save the settings for the XY positions under the phase tab as previously demonstrated.Click the time tab and check the box to select the acquisition interval, frequency and imaging duration. Then click the wave length tab and check the box to select channels to be used in imaging and channel interval frequency. When fished with the settings, click Run Now to initiate the time lapse imaging.Compare to the P.aeruginosa cells in monoculture that remain grouped in rafts when in co-culture with S.aureus, P.aeruginosa has increased single cell motility towards S.aureus colonies. P.aeruginosa single cells will surround and eventually invade S.aureus colonies. Too much or too little humidity during the experiment and incorrectly dried pads are two common problems that lead to drift due to the pad shifting and dragging the cells out of the field of view.Photo bleaching and photo toxicity can cause cells to first, loose their fluorescence then stop dividing and eventually die in subsequent frames. Cells that are initially too close in proximity may not have sufficient time to establish a gradient of secreted signals to which other species can respond. Bacterial cell viability can be visualized by adding propidium iodide to the agarose pads.Green cells indicate live cells actively expressing GFP, while red cells indicate dead propidium iodide stained cells after being treated with P.aeruginosa cell free supernatant. The movement of individual P.aeruginosa cells are tracked from the frame in which a cell leaves the raft through the frame in which the cells reaches the S.aureus cluster. The distance between the P.aeruginosa raft and the S.aureus cluster, provide the Euclidean distance, while the total track lengths provide the accumulated distance.Directedness of each cell is calculated as a ratio of the euclidean distance to the accumulated distance. Wild type P.aeruginosa moves toward wild type S.aureus with significantly higher directedness than toward S.Aureus lacking agr-regulated secreted factors which are necessary for directional motility. When attempting this protocol, it is important to keep in mind that each investigator will need to optimize conditions for drying agarose pads in live imaging based on geographical location, laboratory environment and their particular experiment.This technique reveal interactions between Pseudomonas aeruginosa and Staphylococcus aureus, previously obscured during our investigations of their behavior in bulk culture, bringing us closer to understanding how these organisms interact during infections.

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Research

JoVE Journal

Immunology and Infection

Saliva, Salivary Gland, and Hemolymph Collection from Ixodes scapularis Ticks

Ticks are found worldwide and afflict humans with many tick-borne illnesses. Ticks are vectors for pathogens that cause Lyme disease and tick-borne relapsing fever (Borrelia spp.), Rocky Mountain Spotted fever (Rickettsia rickettsii), ehrlichiosis (Ehrlichia chaffeensis and E. equi), anaplasmosis (Anaplasma phagocytophilum), encephalitis (tick-borne encephalitis virus), babesiosis (Babesia spp.), Colorado tick fever (Coltivirus), and tularemia (Francisella tularensis) 1-8. To be properly transmitted into the host these infectious agents differentially regulate gene expression, interact with tick proteins, and migrate through the tick 3,9-13. For example, the Lyme disease agent, Borrelia burgdorferi, adapts through differential gene expression to the feast and famine stages of the tick's enzootic cycle 14,15. Furthermore, as an Ixodes tick consumes a bloodmeal Borrelia replicate and migrate from the midgut into the hemocoel, where they travel to the salivary glands and are transmitted into the host with the expelled saliva 9,16-19.
As a tick feeds the host typically responds with a strong hemostatic and innate immune response 11,13,20-22. Despite these host responses, I. scapularis can feed for several days because tick saliva contains proteins that are immunomodulatory, lytic agents, anticoagulants, and fibrinolysins to aid the tick feeding 3,11,20,21,23. The immunomodulatory activities possessed by tick saliva or salivary gland extract (SGE) facilitate transmission, proliferation, and dissemination of numerous tick-borne pathogens 3,20,24-27. To further understand how tick-borne infectious agents cause disease it is essential to dissect actively feeding ticks and collect tick saliva. This video protocol demonstrates dissection techniques for the collection of hemolymph and the removal of salivary glands from actively feeding I. scapularis nymphs after 48 and 72 hours post mouse placement. We also demonstrate saliva collection from an adult female I. scapularis tick.

Saliva, Salivary Gland, and Hemolymph Collection from Ixodes scapularis Ticks

The collection of infected tick hemolymph, salivary glands, and saliva is important to study how tick-borne pathogens cause disease. In this protocol we demonstrate how to collect hemolymph and salivary glands from feeding Ixodes scapularis nymphs. We also demonstrate saliva collection from female I. scapularis adults.

Ticks are found worldwide and afflict humans with many tick-borne illnesses. Ticks are vectors for pathogens that cause Lyme disease and tick-borne ...

Visualization of Bacterial Toxin Induced Responses Using Live Cell Fluorescence Microscopy

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

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

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

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 ...

Imaging InlC Secretion to Investigate Cellular Infection by the Bacterial Pathogen Listeria monocytogenes

Bacterial intracellular pathogens can be conceived as molecular tools to dissect cellular signaling cascades due to their capacity to exquisitely manipulate and subvert cell functions which are required for the infection of host target tissues. Among these bacterial pathogens, Listeria monocytogenes is a Gram positive microorganism that has been used as a paradigm for intracellular parasitism in the characterization of cellular immune responses, and which has played instrumental roles in the discovery of molecular pathways controlling cytoskeletal and membrane trafficking dynamics. In this article, we describe a robust microscopical assay for the detection of late cellular infection stages of L. monocytogenes based on the fluorescent labeling of InlC, a secreted bacterial protein which accumulates in the cytoplasm of infected cells; this assay can be coupled to automated high-throughput small interfering RNA screens in order to characterize cellular signaling pathways involved in the up- or down-regulation of infection.

Imaging InlC Secretion to Investigate Cellular Infection by the Bacterial Pathogen Listeria monocytogenes

Listeria monocytogenes is a Gram positive bacterial pathogen frequently used as a major model for the study of intracellular parasitism. Imaging late L. monocytogenes infection stages within the context of small-interfering RNA screens allows for the global study of cellular pathways required for bacterial infection of target host cells.

Bacterial intracellular  pathogens can be conceived as molecular tools to dissect cellular signaling  cascades due to their capacity to exquisitely ...

Visualizing Protein-DNA Interactions in Live Bacterial Cells Using Photoactivated Single-molecule Tracking

Protein-DNA interactions are at the heart of many fundamental cellular processes. For example, DNA replication, transcription, repair, and chromosome organization are governed by DNA-binding proteins that recognize specific DNA structures or sequences. In vitro experiments have helped to generate detailed models for the function of many types of DNA-binding proteins, yet, the exact mechanisms of these processes and their organization in the complex environment of the living cell remain far less understood. We recently introduced a method for quantifying DNA-repair activities in live Escherichia coli cells using Photoactivated Localization Microscopy (PALM) combined with single-molecule tracking. Our general approach identifies individual DNA-binding events by the change in the mobility of a single protein upon association with the chromosome. The fraction of bound molecules provides a direct quantitative measure for the protein activity and abundance of substrates or binding sites at the single-cell level. Here, we describe the concept of the method and demonstrate sample preparation, data acquisition, and data analysis procedures.

Photoactivated localization microscopy (PALM) combined with single-molecule tracking allows direct observation and quantification of protein-DNA interactions in live Escherichia coli cells.

Protein-DNA interactions are at the heart of many fundamental cellular processes. For example, DNA replication, transcription, repair, and chromosome ...

Generation and Multi-phenotypic High-content Screening of Coxiella burnetii Transposon Mutants

Invasion and colonization of host cells by bacterial pathogens depend on the activity of a large number of prokaryotic proteins, defined as virulence factors, which can subvert and manipulate key host functions. The study of host/pathogen interactions is therefore extremely important to understand bacterial infections and develop alternative strategies to counter infectious diseases. This approach however, requires the development of new high-throughput assays for the unbiased, automated identification and characterization of bacterial virulence determinants. Here, we describe a method for the generation of a GFP-tagged mutant library by transposon mutagenesis and the development of high-content screening approaches for the simultaneous identification of multiple transposon-associated phenotypes. Our working model is the intracellular bacterial pathogen Coxiellaburnetii, the etiological agent of the zoonosis Q fever, which is associated with severe outbreaks with a consequent health and economic burden. The obligate intracellular nature of this pathogen has, until recently, severely hampered the identification of bacterial factors involved in host pathogen interactions, making of Coxiella the ideal model for the implementation of high-throughput/high-content approaches.

Generation and Multi-phenotypic High-content Screening of Coxiella burnetii Transposon Mutants

Coxiella burnetii is an obligate intracellular Gram-negative bacterium responsible for the zoonotic disease Q fever. Here we describe methods for the generation of Coxiella fluorescent transposon mutants as well as the automated identification and analysis of the resulting internalization, replication and cytotoxic phenotypes.

Invasion and colonization of host cells by bacterial pathogens depend on the activity of a large number of prokaryotic proteins, defined as virulence ...

Evaluation of the Efficacy And Toxicity of RNAs Targeting HIV-1 Production for Use in Gene or Drug Therapy

Small RNA therapies targeting post-integration steps in the HIV-1 replication cycle are among the top candidates for gene therapy and have the potential to be used as drug therapies for HIV-1 infection. Post-integration inhibitors include ribozymes, short hairpin (sh) RNAs, small interfering (si) RNAs, U1 interference (U1i) RNAs and RNA aptamers. Many of these have been identified using transient co-transfection assays with an HIV-1 expression plasmid and some have advanced to clinical trials. In addition to measures of efficacy, small RNAs have been evaluated for their potential to affect the expression of human RNAs, alter cell growth and/or differentiation, and elicit innate immune responses. In the protocols described here, a set of transient transfection assays designed to evaluate the efficacy and toxicity of RNA molecules targeting post-integration steps in the HIV-1 replication cycle are described. We have used these assays to identify new ribozymes and optimize the format of shRNAs and siRNAs targeting HIV-1 RNA. The methods provide a quick set of assays that are useful for screening new anti-HIV-1 RNAs and could be adapted to screen other post-integration inhibitors of HIV-1 replication.

Methods to evaluate the efficacy and toxicity of RNA molecules targeting post-integration steps of the HIV-1 replication cycle are described. These methods are useful for screening new molecules and optimizing the format of existing ones.

Small RNA therapies targeting post-integration steps in the HIV-1 replication cycle are among the top candidates for gene therapy and have the potential ...

A New Method for Qualitative Multi-scale Analysis of Bacterial Biofilms on Filamentous Fungal Colonies Using Confocal and Electron Microscopy

Bacterial biofilms frequently form on fungal surfaces and can be involved in numerous bacterial-fungal interaction processes, such as metabolic cooperation, competition, or predation. The study of biofilms is important in many biological fields, including environmental science, food production, and medicine. However, few studies have focused on such bacterial biofilms, partially due to the difficulty of investigating them. Most of the methods for qualitative and quantitative biofilm analyses described in the literature are only suitable for biofilms forming on abiotic surfaces or on homogeneous and thin biotic surfaces, such as a monolayer of epithelial cells.
While laser scanning confocal microscopy (LSCM) is often used to analyze in situ and in vivo biofilms, this technology becomes very challenging when applied to bacterial biofilms on fungal hyphae, due to the thickness and the three dimensions of the hyphal networks. To overcome this shortcoming, we developed a protocol combining microscopy with a method to limit the accumulation of hyphal layers in fungal colonies. Using this method, we were able to investigate the development of bacterial biofilms on fungal hyphae at multiple scales using both LSCM and scanning electron microscopy (SEM). This report describes the protocol, including microorganism cultures, bacterial biofilm formation conditions, biofilm staining, and LSCM and SEM visualizations.

This protocol describes a new method to grow and qualitatively analyze bacterial biofilms on fungal hyphae by confocal and electron microscopy.

Bacterial biofilms frequently form on fungal surfaces and can be involved in numerous bacterial-fungal interaction processes, such as metabolic ...

Quantification of Efferocytosis by Single-cell Fluorescence Microscopy

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

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

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

Isolation of Single Intracellular Bacterial Communities Generated from a Murine Model of Urinary Tract Infection for Downstream Single-cell Analysis

In this article, we outline a procedure used to isolate individual intracellular bacterial communities from a mouse that has been experimentally infected in the urinary tract. The protocol can be broadly divided into three sections: the infection, bladder epithelial cell harvesting, and mouth micropipetting to isolate individual infected epithelial cells. The isolated epithelial cell contains viable bacterial cells and is nearly free of contaminating extracellular bacteria, making it ideal for downstream single-cell analysis. The time taken from the start of infection to obtaining a single intracellular bacterial community is about 8 h. This protocol is inexpensive to deploy and uses widely available materials, and we anticipate that it can also be utilized in other infection models to isolate single infected cells from cell mixtures even if those infected cells are rare. However, due to a potential risk in mouth micropipetting, this procedure is not recommended for highly infectious agents.

This protocol describes a simple method of isolating single, infected-bladder epithelial cells from a murine model of urinary tract infection.

In this article, we outline a procedure used to isolate individual intracellular bacterial communities from a mouse that has been experimentally infected ...