This work was supported by the Singapore National Research Foundation (grant NRF-RF2010-06 to J.Y.Y.).

1. Sample Preparation
<ol>
	<li>Cross the Gal43,22 driver line to UAS-GCaMP5G19 flies (w1118;P{20XUAS-IVS-GCaMP5G}attP40). Allow the cross to grow at 25 &#176;C. 
	Note: Generating flies with multiple copies of the Gal4 or UAS-GCaMP transgenes can help to enhance the fluorescent signal intensity. An earlier version of the UAS-GCaMP transgene (UAS-GCaMP3) showed very weak fluorescence intensity when expressed under the control of the Gr68a-Gal4 driver.
	<ol>
		<li>Collect and separate by sex newly eclosed adult flies and raise at 25 &#176;C. 
		Note: The baseline level of GCaMP5G florescence is optimal in flies aged between 14-28 days. Isolating flies by sex precludes the possibility of social interactions that may influence the neural responses.</li>
	</ol>
	</li>
	<li>Anesthetize a fly under a mild stream of CO2.</li>
	<li>Attach the fly to a glass coverslip by first placing a small drop of nail polish in the center of a 0.17 mm coverslip. Gently place a fly on its side on the drop of clear nail polish and ensure that the entire drop is covered by the fly.</li>
	<li>Perform steps 1.5 and 1.6 under a dissecting stereomicroscope, used at 8X magnification.</li>
	<li>Use a wet paintbrush to extend a foreleg of the fly. Secure the foreleg in position by placing two thin strips of tape over the first and fifth tarsal segments (Figure 2A). This will expose tarsal segments T2-T4 for imaging (Figure 2B).</li>
	<li>To image neurons on the proboscis, place a thin strip of tape over the rostrum of the proboscis to expose the labellum (Figure 2C, D).</li>
	<li>Draw a hydrophobic barrier around the fly with a PAP pen to prevent solution from flowing over the coverslip surface. Make measurements within 30 min of mounting.</li>
</ol>
2. Preparation of Stimulus Solution
<ol>
	<li>Dilute lipophilic ligands (such as CH503, used in this study) in a PBST solution: 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4, 0.1% Triton X-100 (v/v); pH 7.4.
	<ol>
		<li>Make a 1 mg/ml stock solution of the ligand using PBST. Prepare all subsequent dilutions of the stock solution using PBST. 
		Note: Depending on the solubility of the ligand of interest, different solvents may be needed to dissolve the ligand. The solubility of CH503 in various solvents is described in Table 1. Unlike other solvents, PBST did not produce an increase in fluorescence.</li>
	</ol>
	</li>
</ol>
3. Stimulation of Gustatory Neurons and Image Acquisition
<ol>
	<li>Using a 10 &#181;l pipettor, place 10 &#181;l of PBST onto the tarsal segments. 
	Note: This step moistens the leg and prevents the tarsal segments from drifting.</li>
	<li>Use an optical system that has a camera of sufficient speed and sensitivity to achieve good fluorescence signal at high time resolution. 
	Note: For this study we used a spinning disk confocal microscope and a sCMOS camera (see Materials List).</li>
	<li>Choose the specific tarsal segment and neurons to be imaged. Acquire three pre-stimulation confocal Z-stacks.
	<ol>
		<li>Ideally, use acquisition settings that are fast enough to allow maximal time resolution while still capturing the entire cell volume. Example acquisition settings are: 200 msec exposure, binning 2 x 2 and 6 x 0.5 &#181;m2 optical sections, captured every 2 sec. 
		Note: Immobilized legs have been imaged successfully for up to 10 min, at 30 sec intervals, with no discernible bleaching of the GCaMP signal. For longer recording times, ensure that the legs are held very&#160;firmly in place on the coverslip.</li>
		<li>Optimize laser power for highest signal-to-noise achievable with minimal photobleaching during the entire imaging regime. For the experiments described here, use a 491 nm diode laser (100 mW) at 30% transmission for imaging both Gr68a and ppk23 gustatory neurons on the legs and proboscis. Use 2 x 2 binning to allow for shorter exposure times while still achieving sufficient spatial resolution. 
		Note: The laser power settings were optimized to allow detection of fluorescent change ranging from a 1.2- to 4.5-fold increase.</li>
	</ol>
	</li>
	<li>Pipette 10 &#181;l of the stimulus solution onto the tarsal segments. For control experiments, add another 10 &#181;l of PBST. Immediately acquire 117 post-stimulation images. 
	Note: The number of post-stimulation images acquired is based on the amount of time in which maximal change in fluorescence was reached (approximately 2 min).
	<ol>
		<li>CRITICAL STEP: Whilst pipetting solution on the preparation, do not to touch the fly with the pipette tip since this will cause the foreleg to move out of position. 
		Note: Alternatively, use a micromanipulator to accurately position a glass capillary containing stimulus close to the tarsal segment to be imaged. Deliver controlled volumes of stimulus using an intracellular microinjection dispenser according to manufacturer&#8217;s instructions.</li>
	</ol>
	</li>
</ol>
4. Image Processing and Analysis: Quantification of the Response
<ol>
	<li>Open a series of confocal Z-stacks using Fiji (<a href="http://fiji.sc/Fiji" target="_blank">http://fiji.sc/Fiji</a>)23 or ImageJ (<a href="http://imagej.nih.gov/ij/" target="_blank">http://imagej.nih.gov/ij/</a>)24 image analysis and processing software.</li>
	<li>Make a sum-intensity projection of all six Z slices for each of the 120 time points.
	<ol>
		<li>Select the &#8220;Stack&#8221; option under &#8220;Image&#8221;. Further, select the &#8220;Z projection&#8221; option and enter &#8220;1&#8221; as the start slice and &#8220;4&#8221; as stop slice. For projection type, select &#8220;sum slices&#8221; from the pull down menu, and choose &#34;All time frames.&#34;</li>
	</ol>
	</li>
	<li>Define the region of interest (ROI) by using the freehand selection tool to draw a boundary around a cell body or neuronal projection. Click on the &#8220;Analyze&#8221; label and select the &#8220;ROI manager&#8221; option under &#8220;Tools&#8221;.</li>
	<li>Quantify the fluorescence of the ROI by selecting the &#8220;Set measurements&#8221; option, under the &#8220;Analyze&#8221; menu and checking the boxes for integrated density, area, mean and maximum gray value and label. Then, select the &#8220;More&#8221; option under the ROI manager menu, and further select the &#8220;Multi measure&#8221; option. 
	Note: This will generate a popup box with values for each of the selected parameters.</li>
	<li>Check the maximum grey values to ensure that none of the pixels have reached saturation (e.g., 65,535 for a 16-bit camera). 
	NOTE: For the experiments shown in this paper, responses were measured either from cell bodies or neuronal projections, depending on where the maximal increase in &#916;F/F was observed.</li>
	<li>Calculate the maximum relative change in fluorescence (&#916;F/F) at time point t is as follows: 
	<img alt="Equation 1" src="/files/ftp_upload/53392/53392eq1.jpg" />
	<ol>
		<li>Subtract background fluorescence from each F(cell) value. To quantify background fluorescence, measure the integrated density of a ROI of equivalent size and shape from an area of the foreleg (or proboscis) devoid of detectable neurons.</li>
		<li>To quantify the post-stimulation fluorescence, F(cell)t, measure the integrated density of the ROI of the cell body or process at time point &#8220;t&#8221; after stimulation.</li>
		<li>To quantify the pre-stimulation fluorescence, F(cell)t=0, analyze pre-stimulus images of the cell body or process in the same way as the post-stimulation signal. Use the mean integrated density of 3 images to determine F(cell)t=0.</li>
	</ol>
	</li>
</ol>

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The authors declare that there are no competing financial interests.

We describe here a method to perform live calcium imaging of Drosophila peripheral neurons in 2 different sensory organs. The Ca2+-evoked GCaMP fluorescent responses in Gr68a-neurons induced by the pheromone ligand CH503 were dose-dependent and quantitative. It was also possible to discern different neural response patterns such as phasic and tonic responses.
Neurons showing phasic responses are believed to allow rapid adaptation to continuous stimuli. This type of response...

Animals rely on olfactory and gustatory information to mediate decisions essential for survival and reproduction. Understanding how chemosensory cues are detected and processed by the nervous system requires identification of the sensory receptor(s) and the corresponding chemical ligands. Drosophila detect a staggering variety of volatile and non-volatile compounds and are an excellent model in which to study the physiological mechanisms underlying chemosensation. While the olfactory organs perceive volatile molecules, the gustatory organs are specialized to detect low volatility compounds. Here, we present a method to directly measure neuronal responses from the taste organs of Drosophila melanogaster to low-volatility, lipophilic ligands.
Gustatory organs of the fly include the forelegs, proboscis, and wings. Distributed on the surface of the taste organs are hair-like structures known as sensilla that respond to sugars, bitters, salts, water and pheromones8. The sensilla have been classified morphologically into taste bristles and taste pegs9. There are about 31 taste bristles on the labellum that are classified into the long (l-type), short (s-type) and intermediate (i-type) morphologies. The &#8216;l&#8217; and &#8216;s&#8217; sensilla house 4 sensory neurons that respond physiologically to sugar (the S cell), low salt (the L1 cell), high salt and bitter compounds (the L2 cell) and water (the W cell)10,11. The &#8216;i&#8217; sensilla house 2 sensory neurons, one of which responds both to low salt and sugar, while the other responds to high salt12. There are approximately 41 taste sensilla in males and 26 sensilla in females distributed on each of the forelegs. For both males and females, there are 21 sensilla on the midleg, and about 22 sensilla on the hindleg13. The gustatory neurons enclosed by the taste sensilla on the legs are also classified into L1, L2, W and S types.
One standard method for measuring electrical activity from single neurons uses extracellular or intracellular electrodes to record ion flow. Electrode measurements allow neuronal function to be studied in non-model organisms such as Drosophila species, moths, and bees which lack extensive genetic tools for neural labeling. However, while electrophysiological methods have been routinely used to measure activity from Drosophila taste sensilla4,13,14, applying this approach presents several technical challenges. First, taste bristles need to be identified based on morphology and spatial location. Upon identification, electrophysiological measurements can be hindered by the small size of the sensilla, limited accessibility due to position, and difficulty in applying controlled volumes of a chemical stimulus to a taste bristle. Also, stimulation of sensilla may generate signals from more than one neuronal type15. Second, detection of electrical signals can be confounded by background noise resulting from mechanical vibrations and noise from electronic equipment. Third, the use of a sharpened electrode can damage the fly preparation, if used incorrectly14. Finally, assembling an electrophysiology rig requires specialized electronic components for stimulus delivery, signal recording, and data analysis and can be costly.
In D. melanogaster, the availability of genetically-encoded tools has facilitated the development of imaging techniques that allow the responses of small populations of neurons to be studied. One such approach is the use of the CaLexA reporter16. In this method, gene sequences encoding the LexA-VP16 transcription factor and a calcium sensitive protein NFAT are fused together. Calcium activation of the protein phosphatase calcineurin catalyzes the de-phosphorylation of NFAT and, in turn, facilitates its import into the nucleus. Inside the nucleus, the LexA domain binds to a LexAop-DNA binding motif which directs the expression of a green fluorescent protein (GFP) reporter, thus allowing sustained identification of functionally activated neurons. The approach has been used successfully to measure the response of specific olfactory glomeruli in the antennal lobe following exposure of live flies to an odorant16. Recently, physiological responses of IR52c gustatory receptor neurons were measured from live flies using CaLexA7. However, for this study, it was necessary to age flies for up to 6 weeks to enhance GFP transcription. While immunostaining with an anti-GFP antibody can be used to boost the CaLexA signal, this method would require tissue fixation, thus precluding possibility for live-cell imaging.
The genetically encoded calcium indicator protein GCaMP has also been extensively used to study neuronal responses in a number of species17. The protein GCaMP fluoresces with low intensity prior to neuronal stimulation. Application of a stimulus triggers an action potential in the neuron, resulting in the influx of Ca2+. GCaMP, bound to Ca2+, undergoes a conformational change, causing it to fluoresce with brighter intensity (Figure 1). A recently developed single&#8211;neuron calcium imaging approach was used to identify glucose as the ligand for the foreleg specific Gr61a gustatory neurons18. In this study, dissected forelegs from transgenic Drosophila expressing GCaMP in Gr61a gustatory neurons were covered with a layer of agarose prior to imaging. However, the use of dissected tissue can cause eventual rundown of GCaMP expression, thus limiting measurement time and detection sensitivity. In addition, agarose can limit sensitivity due to high background levels of fluorescence and its light scattering properties.
To address some of these drawbacks, we describe the use of GCaMP-mediated calcium imaging to record physiological responses from foreleg and proboscis gustatory neurons of intact animals. We show the physiological responses of Gr68a and ppk23 neurons expressing genetically encoded GCaMP5G19 to a Drosophila lipid pheromone, (3R, 11Z, 19Z)-3-acetoxy-11,19-octacosadien-1-ol (CH503)20,21. Neuronal responses are measured by quantifying the increase in fluorescence of the GCaMP5G signal during pheromone stimulation. In this protocol, neurons are imaged for a total duration of 120 sec, which is sufficient to differentiate patterns of neuronal activation in individual cells.

<table>
	<tbody>
		<tr>
			<td>Gr68a-Gal4</td>
			<td></td>
			<td></td>
			<td>Gift from H. Amrein (Texas A&#38;M Health Science Center, TX, USA) and J. Carlson (Yale University, CT, USA)</td>
		</tr>
		<tr>
			<td>ppk23-Gal4</td>
			<td></td>
			<td></td>
			<td>Gift from K. Scott (Univ. of California, Berkeley, CA, USA)</td>
		</tr>
		<tr>
			<td>UAS-GCaMP5&#160;</td>
			<td></td>
			<td>42037</td>
			<td>Bloomington Drosophila&#160; Stock Center</td>
		</tr>
		<tr>
			<td>0.17 mm coverslip (Gold-Seal coverslip)</td>
			<td>Electron Microscopy Services</td>
			<td>63790-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Nail polish, &#34;Hard as Nails Clear&#34;&#160;</td>
			<td>Sally Hansen</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PAP pen</td>
			<td>Sigma-Aldrich&#160;</td>
			<td>Z377821</td>
			<td></td>
		</tr>
		<tr>
			<td>Paint brush</td>
			<td></td>
			<td></td>
			<td>fine-tipped brush</td>
		</tr>
		<tr>
			<td>Tape&#160;</td>
			<td>Scotch brand</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma-Aldrich&#160;</td>
			<td>13021</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol, lab grade</td>
			<td>Merck</td>
			<td>10094</td>
			<td></td>
		</tr>
		<tr>
			<td>Hexane, HPLC grade</td>
			<td>Sigma-Aldrich&#160;</td>
			<td>H303SK-4</td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Sigma-Aldrich&#160;</td>
			<td>472301</td>
			<td></td>
		</tr>
		<tr>
			<td>PBST</td>
			<td></td>
			<td></td>
			<td>Recipe described in the protocol section</td>
		</tr>
		<tr>
			<td>CH503</td>
			<td></td>
			<td></td>
			<td>Synthesis described in Mori et al., 2010</td>
		</tr>
		<tr>
			<td>sCMOS Camera (ORCA Flash4.0)</td>
			<td>Hamamatsu&#160;</td>
			<td>C11578-22U</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope (Ti-Eclipse)</td>
			<td>Nikon</td>
			<td>Ni-E</td>
			<td></td>
		</tr>
		<tr>
			<td>Spinning Disk Scan head&#160;</td>
			<td>Yokogawa</td>
			<td>CSU-X1-A1</td>
			<td></td>
		</tr>
		<tr>
			<td>Aquistion Software (MetaMorph Premier)</td>
			<td>Molecular Devices</td>
			<td>40002</td>
			<td></td>
		</tr>
		<tr>
			<td>Fiji software</td>
			<td>open source</td>
			<td></td>
			<td><a href="http://fiji.sc/Fiji">http://fiji.sc/Fiji</a></td>
		</tr>
	</tbody>
</table>

measuring physiological responses of drosophila sensory neurons to lipid pheromones using live calcium imaging

Unlike mammals, insects such as Drosophila have multiple taste organs. The chemosensory neurons on the legs, proboscis, wings and ovipositor of Drosophila express gustatory receptors1,2, ion channels3-6, and ionotropic receptors7 that are involved in the detection of volatile and non-volatile sensory cues. These neurons directly contact tastants such as food, noxious substances and pheromones and therefore influence many complex behaviors such as feeding, egg-laying and mating. Electrode recordings and calcium imaging have been widely used in insects to quantify the neuronal responses evoked by these tastants. However, electrophysiology requires specialized equipment and obtaining measurements from a single taste sensillum can be technically challenging depending on the cell-type, size, and position. In addition, single neuron resolution in Drosophila can be difficult to achieve since taste sensilla house more than one type of chemosensory neuron. The live calcium imaging method described here allows responses of single gustatory neurons in live flies to be measured. This method is especially suitable for imaging neuronal responses to lipid pheromones and other ligand types that have low solubility in water-based solvents.

The GCaMP calcium indicator was genetically expressed using Gr68a-Gal4 or ppk23-Gal4 drivers. Distinct populations of foreleg neurons and non-neural support cells are labeled by each driver (Figure 3A-C). Ligand-specific responses to the lipophilic pheromone CH503 were observed in Gr68a-Gal4 and ppk23-Gal4 cells expressing GCaMP (Figure 3D). The relative change in fluorescence intensity of the GCaMP5G signal (&#916;F/F) and increased with high...

Watch this Scientific Journal Video about Measuring Physiological Responses of Drosophila Sensory Neurons to Lipid Pheromones Using Live Calcium Imaging at JoVE.com

Measuring Physiological Responses of Drosophila Sensory Neurons to Lipid Pheromones Using Live Calcium Imaging

The forelegs and proboscis of Drosophila contain a rich repertoire of gustatory sensory neurons. Here, we present a method using calcium imaging to measure physiological responses from sensory neurons in the foreleg and proboscis of live flies upon exogenous application of a gustatory pheromone.

Measuring Physiological Responses of Drosophila Sensory Neurons to Lipid Pheromones Using Live Calcium Imaging

Unlike mammals, insects such as Drosophila have multiple taste organs. The chemosensory neurons on the legs, proboscis, wings and ovipositor of Drosophila ...

measuring-physiological-responses-drosophila-sensory-neurons-to-lipid

Research

JoVE Journal

Immunology and Infection

8.5K Views.  Temasek Life Sciences Laboratory. The forelegs and proboscis of Drosophila contain a rich repertoire of gustatory sensory neurons. Here, we present a method using calcium imaging to measure physiological responses from sensory neurons in the foreleg and proboscis of live flies upon exogenous application of a gustatory pheromone. 

Video: Measuring Physiological Responses of Drosophila Sensory Neurons to Lipid Pheromones Using Live Calcium Imaging

Assessing Stomatal Response to Live Bacterial Cells using Whole Leaf Imaging

Stomata are natural openings in the plant epidermis responsible for gas exchange between plant interior and environment. They are formed by a pair of guard cells, which are able to close the stomatal pore in response to a number of external factors including light intensity, carbon dioxide concentration, and relative humidity (RH). The stomatal pore is also the main route for pathogen entry into leaves, a crucial step for disease development. Recent studies have unveiled that closure of the pore is effective in minimizing bacterial disease development in Arabidopsis plants; an integral part of plant innate immunity. Previously, we have used epidermal peels to assess stomatal response to live bacteria (Melotto et al. 2006); however maintaining favorable environmental conditions for both plant epidermal peels and bacterial cells has been challenging. Leaf epidermis can be kept alive and healthy with MES buffer (10 mM KCl, 25 mM MES-KOH, pH 6.15) for electrophysiological experiments of guard cells. However, this buffer is not appropriate for obtaining bacterial suspension. On the other hand, bacterial cells can be kept alive in water which is not proper to maintain epidermal peels for long period of times. When an epidermal peel floats on water, the cells in the peel that are exposed to air dry within 4 hours limiting the timing to conduct the experiment. An ideal method for assessing the effect of a particular stimulus on guard cells should present minimal interference to stomatal physiology and to the natural environment of the plant as much as possible. We, therefore, developed a new method to assess stomatal response to live bacteria in which leaf wounding and manipulation is greatly minimized aiming to provide an easily reproducible and reliable stomatal assay. The protocol is based on staining of intact leaf with propidium iodide (PI), incubation of staining leaf with bacterial suspension, and observation of leaves under laser scanning confocal microscope. Finally, this method allows for the observation of the same live leaf sample over extended periods of time using conditions that closely mimic the natural conditions under which plants are attacked by pathogens.

We have developed a simple and reproducible protocol to access stomatal response to live bacteria. This method minimizes wounding and manipulation of the leaf as compared to the use of epidermal peels reported previously.

Stomata are natural openings in the plant epidermis responsible for gas exchange between plant interior and environment. They are formed by a pair of ...

Measuring Bacterial Load and Immune Responses in Mice Infected with Listeria monocytogenes

Listeria monocytogenes (Listeria) is a Gram-positive facultative intracellular pathogen1. Mouse studies typically employ intravenous injection of Listeria, which results in systemic infection2. After injection, Listeria quickly disseminates to the spleen and liver due to uptake by CD8&alpha;+ dendritic cells and Kupffer cells3,4. Once phagocytosed, various bacterial proteins enable Listeria to escape the phagosome, survive within the cytosol, and infect neighboring cells5. During the first three days of infection, different innate immune cells (e.g. monocytes, neutrophils, NK cells, dendritic cells) mediate bactericidal mechanisms that minimize Listeria proliferation. CD8+ T cells are subsequently recruited and responsible for the eventual clearance of Listeria from the host, typically within 10 days of infection6.
Successful clearance of Listeria from infected mice depends on the appropriate onset of host immune responses6	. There is a broad range of sensitivities amongst inbred mouse strains7,8. Generally, mice with increased susceptibility to Listeria infection are less able to control bacterial proliferation, demonstrating increased bacterial load and/or delayed clearance compared to resistant mice. Genetic studies, including linkage analyses and knockout mouse strains, have identified various genes for which sequence variation affects host responses to Listeria infection6,8-14. Determination and comparison of infection kinetics between different mouse strains is therefore an important method for identifying host genetic factors that contribute to immune responses against Listeria. Comparison of host responses to different Listeria strains is also an effective way to identify bacterial virulence factors that may serve as potential targets for antibiotic therapy or vaccine design.
We describe here a straightforward method for measuring bacterial load (colony forming units [CFU] per tissue) and preparing single-cell suspensions of the liver and spleen for FACS analysis of immune responses in Listeria-infected mice. This method is particularly useful for initial characterization of Listeria infection in novel mouse strains, as well as comparison of immune responses between different mouse strains infected with Listeria. We use the Listeria monocytogenes EGD strain15 that, when cultured on blood agar, exhibits a characteristic halo zone around each colony due to &beta;-hemolysis1 (Figure 1). Bacterial load and immune responses can be determined at any time-point after infection by culturing tissue homogenate on blood agar plates and preparing tissue cell suspensions for FACS analysis using the protocols described below. We would note that individuals who are immunocompromised or pregnant should not handle Listeria, and the relevant institutional biosafety committee and animal facility management should be consulted before work commences.

Measuring Bacterial Load and Immune Responses in Mice Infected with Listeria monocytogenes

Listeria monocytogenes is a model organism for studying immune responses and genetic susceptibility to intracellular bacteria in mice. This method enables one to measure bacterial load and generate single-cell suspensions of the liver and spleen from mice for FACS analysis to determine changes in immune cells due to Listeria infection.

Listeria monocytogenes (Listeria) is a Gram-positive facultative intracellular pathogen1. Mouse studies typically employ intravenous injection of ...

Live Cell Imaging of Bacillus subtilis and Streptococcus pneumoniae using Automated Time-lapse Microscopy

During the last few years scientists became increasingly aware that average data obtained from microbial population based experiments are not representative of the behavior, status or phenotype of single cells. Due to this new insight the number of single cell studies rises continuously (for recent reviews see 1,2,3). However, many of the single cell techniques applied do not allow monitoring the development and behavior of one specific single cell in time (e.g. flow cytometry or standard microscopy).
Here, we provide a detailed description of a microscopy method used in several recent studies 4, 5, 6, 7, which allows following and recording (fluorescence of) individual bacterial cells of Bacillus subtilis and Streptococcus pneumoniae through growth and division for many generations. The resulting movies can be used to construct phylogenetic lineage trees by tracing back the history of a single cell within a population that originated from one common ancestor. This time-lapse fluorescence microscopy method cannot only be used to investigate growth, division and differentiation of individual cells, but also to analyze the effect of cell history and ancestry on specific cellular behavior. Furthermore, time-lapse microscopy is ideally suited to examine gene expression dynamics and protein localization during the bacterial cell cycle. The method explains how to prepare the bacterial cells and construct the microscope slide to enable the outgrowth of single cells into a microcolony. In short, single cells are spotted on a semi-solid surface consisting of growth medium supplemented with agarose on which they grow and divide under a fluorescence microscope within a temperature controlled environmental chamber. Images are captured at specific intervals and are later analyzed using the open source software ImageJ.

Live Cell Imaging of Bacillus subtilis and Streptococcus pneumoniae using Automated Time-lapse Microscopy

This protocol provides a step-by-step procedure to monitor single cell behavior of different bacteria in time using automated fluorescence time-lapse microscopy. Furthermore, we provide guidelines how to analyze the microscopy images.

During the last few years scientists became increasingly aware that average data obtained from microbial population based experiments are not ...

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 ...

Live Cell Imaging of Alphaherpes Virus Anterograde Transport and Spread

Advances in live cell fluorescence microscopy techniques, as well as the construction of recombinant viral strains that express fluorescent fusion proteins have enabled real-time visualization of transport and spread of alphaherpes virus infection of neurons. The utility of novel fluorescent fusion proteins to viral membrane, tegument, and capsids, in conjunction with live cell imaging, identified viral particle assemblies undergoing transport within axons. Similar tools have been successfully employed for analyses of cell-cell spread of viral particles to quantify the number and diversity of virions transmitted between cells. Importantly, the techniques of live cell imaging of anterograde transport and spread produce a wealth of information including particle transport velocities, distributions of particles, and temporal analyses of protein localization. Alongside classical viral genetic techniques, these methodologies have provided critical insights into important mechanistic questions. In this article we describe in detail the imaging methods that were developed to answer basic questions of alphaherpes virus transport and spread.

Live cell imaging of alphaherpes virus infections enables analysis of the dynamic events of directed transport and intercellular spread. Here, we present methodologies that utilize recombinant viral strains expressing fluorescent fusion proteins to facilitate visualization of viral assemblies during infection of primary neurons.

Advances in live cell fluorescence microscopy  techniques, as well as the construction of recombinant viral strains that  express fluorescent fusion ...

Phage Phenomics: Physiological Approaches to Characterize Novel Viral Proteins

Current investigations into phage-host interactions are dependent on extrapolating knowledge from (meta)genomes. Interestingly, 60 - 95% of all phage sequences share no homology to current annotated proteins. As a result, a large proportion of phage genes are annotated as hypothetical. This reality heavily affects the annotation of both structural and auxiliary metabolic genes. Here we present phenomic methods designed to capture the physiological response(s) of a selected host during expression of one of these unknown phage genes. Multi-phenotype Assay Plates (MAPs) are used to monitor the diversity of host substrate utilization and subsequent biomass formation, while metabolomics provides bi-product analysis by monitoring metabolite abundance and diversity. Both tools are used simultaneously to provide a phenotypic profile associated with expression of a single putative phage open reading frame (ORF). Representative results for both methods are compared, highlighting the phenotypic profile differences of a host carrying either putative structural or metabolic phage genes. In addition, the visualization techniques and high throughput computational pipelines that facilitated experimental analysis are presented.

Here, we present phenomic approaches for the functional characterization of putative phage genes. Techniques include a developed assay capable of monitoring host anabolic metabolism, the Multi-phenotype Assay Plates (MAPs), in addition to the established method of metabolomics, capable of measuring effects to catabolic metabolism.

Current investigations into phage-host interactions are dependent on extrapolating knowledge from (meta)genomes. Interestingly, 60 - 95% of all phage ...

An In Vitro Model for Measuring Immune Responses to Malaria in the Context of HIV Co-infection

Malaria and HIV co-infection is a growing health priority. However, most research on malaria or HIV currently focuses on each infection individually. Although understanding the disease dynamics for each of these pathogens independently is vital, it is also important that the interactions between these pathogens are investigated and understood. 
We have developed a versatile in vitro model of HIV-malaria co-infection to study host immune responses to malaria in the context of HIV infection. Our model allows the study of secreted factors in cellular supernatants, cell surface and intracellular protein markers, as well as RNA expression levels. The experimental design and methods used limit variability and promote data reliability and reproducibility. 
All pathogens used in this model are natural human pathogens (Plasmodium falciparum and HIV-1), and all infected cells are naturally infected and used fresh. We use human erythrocytes parasitized with P. falciparum and maintained in continuous in vitro culture. We obtain freshly isolated peripheral blood mononuclear cells from chronically HIV-infected volunteers. Every condition used has an appropriate control (P. falciparum parasitized vs. normal erythrocytes), and every HIV-infected donor has an HIV uninfected control, from which cells are harvested on the same day. This model provides a realistic environment to study the interactions between malaria parasites and human immune cells in the context of HIV infection.

An In Vitro Model for Measuring Immune Responses to Malaria in the Context of HIV Co-infection

Human co-infection is difficult to replicate in vitro. However, human malaria parasites can readily be cultured in vitro, as can freshly isolated human peripheral blood mononuclear cells naturally infected with HIV. This provides an excellent model for studying early immune responses to malaria parasites in the context of HIV co-infection.

Malaria and HIV co-infection is a growing health priority. However, most research on malaria or HIV currently focuses on each infection individually. ...

Methods to Study Lipid Alterations in Neutrophils and the Subsequent Formation of Neutrophil Extracellular Traps

Lipid analysis performed by high performance thin layer chromatography (HPTLC) is a relatively simple, cost-effective method of analyzing a broad range of lipids. The function of lipids (e.g., in host-pathogen interactions or host entry) has been reported to play a crucial role in cellular processes. Here, we show a method to determine lipid composition, with a focus on the cholesterol level of primary blood-derived neutrophils, by HPTLC in comparison to high performance liquid chromatography (HPLC). The aim was to investigate the role of lipid/cholesterol alterations in the formation of neutrophil extracellular traps (NETs). NET release is known as a host defense mechanism to prevent pathogens from spreading within the host. Therefore, blood-derived human neutrophils were treated with methyl-&#946;-cyclodextrin (M&#946;CD) to induce lipid alterations in the cells. Using HPTLC and HPLC, we have shown that M&#946;CD treatment of the cells leads to lipid alterations associated with a significant reduction in the cholesterol content of the cell. At the same time, M&#946;CD treatment of the neutrophils led to the formation of NETs, as shown by immunofluorescence microscopy. In summary, here we present a detailed method to study lipid alterations in neutrophils and the formation of NETs.

Lipids are known to play an important role in cellular functions. Here, we describe a method to determine the lipid composition of neutrophils, with emphasis on the cholesterol level, by using both HPTLC and HPLC to gain a better understanding of the underlying mechanisms of neutrophil extracellular trap formation.

Lipid analysis performed by high performance thin layer chromatography (HPTLC) is a relatively simple, cost-effective method of analyzing a broad range of ...

Measuring Influenza Neutralizing Antibody Responses to A(H3N2) Viruses in Human Sera by Microneutralization Assays Using MDCK-SIAT1 Cells

Neutralizing antibodies against hemagglutinin (HA) of influenza viruses are considered the main immune mechanism that correlates with protection for influenza infections. Microneutralization (MN) assays are often used to measure neutralizing antibody responses in human sera after influenza vaccination or infection. Madine Darby Canine Kidney (MDCK) cells are the commonly used cell substrate for MN assays. However, currently circulating 3C.2a and 3C.3a A(H3N2) influenza viruses have acquired altered receptor binding specificity. The MDCK-SIAT1 cell line with increased &#945;-2,6 sialic galactose moieties on the surface has proven to provide improved infectivity and more faithful replications than conventional MDCK cells for these contemporary A(H3N2) viruses. Here, we describe a MN assay using MDCK-SIAT1 cells that has been optimized to quantify neutralizing antibody titers to these contemporary A(H3N2) viruses. In this protocol, heat inactivated sera containing neutralizing antibodies are first serially diluted, then incubated with 100 TCID50/well of influenza A(H3N2) viruses to allow antibodies in the sera to bind to the viruses. MDCK-SIAT1 cells are then added to the virus-antibody mixture, and incubated for 18 - 20 h at 37 &#176;C, 5% CO2 to allow A(H3N2) viruses to infect MDCK-SIAT1 cells. After overnight incubation, plates are fixed and the amount of virus in each well is quantified by an enzyme-linked immunosorbent assay (ELISA) using anti-influenza A nucleoprotein (NP) monoclonal antibodies. Neutralizing antibody titer is defined as the reciprocal of the highest serum dilution that provides &#8805;50% inhibition of virus infectivity.

Measuring Influenza Neutralizing Antibody Responses to AH3N2 Viruses in Human Sera by Microneutralization Assays Using MDCK-SIAT1 Cells

Influenza neutralizing antibodies correlate with protection of influenza infections. Microneutralization assays measure neutralizing antibodies in human sera and are often used for influenza human serology. We describe a microneutralization assay using MDCK-SIAT1 cells to measure neutralizing antibody titers to contemporary 3C.2a and 3C.3a A(H3N2) viruses following influenza vaccination or infection.

Neutralizing antibodies against hemagglutinin (HA) of influenza viruses are considered the main immune mechanism that correlates with protection for ...

Imaging Ca2+ Responses During Shigella Infection of Epithelial Cells

Ca2+ is a ubiquitous ion involved in all known cellular processes. While global Ca2+ responses may affect cell fate, local variations in free Ca2+ cytosolic concentrations, linked to release from internal stores or an influx through plasma membrane channels, regulate cortical cell processes. Pathogens that adhere to or invade host cells trigger a reorganization of the actin cytoskeleton underlying the host plasma membrane, which likely affects both global and local Ca2+ signaling. Because these events may occur at low frequencies in a pseudo-stochastic manner over extended kinetics, the analysis of Ca2+ signals induced by pathogens raises major technical challenges that need to be addressed.
Here, we report protocols for the detection of global and local Ca2+ signals upon a Shigella infection of epithelial cells. In these protocols, artefacts linked to a prolonged exposure and photodamage associated with the excitation of Ca2+ fluorescent probes are troubleshot by stringently controlling the acquisition parameters over defined time periods during a Shigella invasion. Procedures are implemented to rigorously analyze the amplitude and frequency of global cytosolic Ca2+ signals during extended infection kinetics using the chemical probe Fluo-4.

Imaging Ca2+ Responses During Shigella Infection of Epithelial Cells

Here, we present protocols to visualize calcium (Ca2+) responses elicited by HeLa cells infected by Shigella. By optimizing the parameters of bacterial infection and imaging with Ca2+ fluorescent probes, atypical global and local Ca2+ signals induced by bacteria over a large range of infection kinetics are characterized.