The authors thank Prof. Sven Krappmann (University Hospital Erlangen and FUA Erlangen-N&#252;rnberg, Germany) for providing the Aspergillus fumigatus conidia strain AfS150. The authors thank MIPT Press Office. V.B. acknowledges the Ministry of Science and Higher Education of the Russian Federation (#075-00337-20-03, project FSMG-2020-0003). The work regarding A. fumigatus conidia imaging and quantification was supported by RSF &#8470; 19-75-00082. The work regarding airways imaging was supported by RFBR &#8470; 20-04-60311.

All methods concerning laboratory animals described here have been approved by the Institutional Animal Care and Use Committee (IACUC) at the Shemyakin and Ovchinnikov Institute of Bioorganic Chemistry, Russian Academy of Sciences (protocol number 226/2017).
1. A. fumigatus conidia application
<ol>
	<li>To obtain fluorescently labeled A. fumigatus conidia, fix 5 &#215; 108 conidia by adding 1 mL of 3% paraformaldehyde to the conidia pellet. Incubate in a 50 mL test tube for 2 h on a shaker at room temperature.</li>
	<li>Wash conidia with 20 mL of phosphate buffer saline (PBS): centrifuge at 1,000 x g for 15 min, gently remove the supernatant, and add&#160;fresh PBS in a volume of 20 mL. Repeat.</li>
	<li>Dissolve the conidia in 900 &#181;L of 0.1 M NaHCO3.</li>
	<li>Dissolve the succinimidyl ester of the 594 nm fluorescent dye in 100 &#181;L of dimethyl sulfoxide and add to conidia.</li>
	<li>Incubate the conidia with the dye for 1 h on a shaker at 150 rpm at room temperature.</li>
	<li>Wash the conidia twice with 20 mL of PBS in the centrifuge at 1,000 x g for 15 min.</li>
	<li>Dilute the conidia to a concentration of 1 &#215; 108 conidia/mL in PBS and store at 4 &#176;C.</li>
	<li>Anesthetize the mouse with 0.5-3% isoflurane vapor. Put the mouse on the holder, fix the tongue with smooth forceps, and hold the nares. Take a single channel pipette and apply 50 &#181;L of conidia suspension to the mouse pharynx. Wait until the suspension is inhaled.</li>
</ol>
2. Specimen preparation
<ol>
	<li>Prepare the 50 mL test tube and fill it with 15 mL of fresh 2% paraformaldehyde. Place the medical instruments (15 cm toothed dissecting forceps, 8 cm fine-tipped forceps, and 10 cm scissors with blunt ends) into 70% ethanol solution.</li>
	<li>Euthanize the mouse in accordance with the IACUC protocol. Then place the mouse in the dorsal position and fix the mouse paws with needles. 
	NOTE: If using cervical dislocation for euthanasia, ensure the integrity of the trachea.
	<ol>
		<li>Treat the mouse with 70% ethanol using a sprayer.</li>
		<li>Make a median longitudinal cut in the ventral skin from the hind paws to the forepaws and the chin.</li>
		<li>Separate the skin from the subcutaneous tissue using toothed forceps and closed scissors. Fix the upper ends of the skin with needles.</li>
		<li>Make an incision in the abdominal wall and separate the liver from the diaphragm. Carefully pick the diaphragm with closed scissors and then cut the diaphragm.</li>
		<li>Make a medial cut of the thorax and neck connective tissues until the trachea is visible.</li>
	</ol>
	</li>
	<li>Run a silk thread beneath the trachea and make a surgical knot using two forceps. 
	NOTE: Alternatively, use dental floss instead of silk thread.
	<ol>
		<li>Carefully pull the thread and cut off the lungs from the connective tissue with scissors. Hold the scissors perpendicular to the table.</li>
		<li>Put the lungs in the 50 mL test tube with 2% paraformaldehyde. Leave thread ends outside the tube, put the cover tight, and turn over the tube to cover the lungs with paraformaldehyde. Hold the lungs overnight at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Dissect the lung lobes from the heart and each other with a scalpel.
	<ol>
		<li>Put the lung lobes in a 24 well plate, with each lobe in a separate well. Wash the lung lobes in 1 mL of Tris-buffered saline (TBS) pH 7.4, 5 times for 1 h each on a shaker at 150 rpm.</li>
		<li>Replace 1 mL of TBS with 1 mL of the blocking buffer (1% Triton X 100, 5% powdered milk in TBS) and leave the specimen overnight at room temperature on a shaker.</li>
		<li>Replace the blocking buffer with 1 mL of streptavidin-488- nm fluorchrome conjugate diluted 1:30 in TBS. Leave the specimen for at least 72 h at room temperature on a shaker (150 rpm).</li>
	</ol>
	</li>
	<li>Wash the specimen 5 times for 1 h each in 1 mL of TBS at room temperature on a shaker (150 rpm). Transfer the specimen to the new wells and cover it with 2% paraformaldehyde overnight at 4 &#176;C for post-fixation.</li>
</ol>
3. Mouse lung lobe optical clearing
<ol>
	<li>Place the specimen in the 5 mL glass bottle filled with 3 mL of 50% methanol-water solution and put it on to the sample mixer at room temperature for 1 h.</li>
	<li>Replace 3 mL of 50% methanol with 3 mL of 100% methanol and put it on to the sample mixer for 2 h. Prepare a 1:2 v/v mixture of benzyl alcohol and benzyl benzoate (BABB) in a total volume of 1 mL.</li>
	<li>Transfer the specimen to a 24 well plate and cover with 1 mL of BABB mixture for at least 30 min. Do not leave the specimen in BABB for a long time; BABB can damage the plastic plate and make the specimen too rigid.</li>
	<li>Put the specimen in the cell imaging coverglass-bottom chamber. The sample is ready for imaging.</li>
</ol>
4. Mouse lung lobe imaging with CLSM
<ol>
	<li>Turn the microscope system on. Open the microscope software. Turn the transmission light on in the Locate tab. Select the 10x objective. 
	NOTE: For more detailed analysis, use the 20x objective, but it increases the time of the experiment and the image file size.</li>
	<li>Put the chamber with the specimen in the cover slide holder. Put the holder on the XY stage over the objective. Center the specimen using the XY controls of the stage on top of the objective. Use the transmission light and eyepiece to manually find the specimen Z-plane.</li>
	<li>Switch the software to the Acquisition tab. Select CLSM &#955;-mode. Turn the 488 nm and 561 nm lasers on. Select the 488/561 nm dichroic mirror. Change the spectral range of the detector to 490-695 nm. Adjust the laser power to the appropriate range (10-50 &#181;W). Adjust the Detector Gain between 750-900 range. 
	NOTE: Higher gain settings are undesirable due to noise.</li>
	<li>Narrow the pinhole to 1 Airy unit by clicking 1 AU button. Set pixel resolution to 512 &#215; 512 pixels.</li>
	<li>Switch on the Z-stack mode. Start Live Imaging. Select the focal plane in which both dyes are visible. 
	NOTE: If necessary, adjust the laser power to normalize the brightness of the two fluorescent dyes. Use Gallery and Single-channel mode, to avoid clipping and to precisely match fluorescence intensity.</li>
	<li>Expand the appeared Z-stack pane. Use the focusing wheel and find the lowest plane of the sample. Use the First button in the Z-stack pane. Move the focal plane upwards to find the top boundary of the specimen and save the position by using the Last button. Check the representation of the sample depth in the Z-stack pane. 
	NOTE: A representation of the sample depth will appear in the Z-stack pane after the First and the Last positions are chosen.</li>
	<li>Position the objective using the focusing wheel near the bottom of the specimen, by looking at the Z-stack pane. This is approximately 20 &#181;m from the bottom of the specimen.</li>
	<li>Turn the Z-stack mode off and turn the Tile Scan mode on. Acquire the image with an appropriate number of tiles for the size of the specimen. 5 &#215; 5 tiles are a good starting point.</li>
	<li>Adjust the number of tiles and the specimen XY position until the whole lung fits inside the tiled image. Recheck the correctness of the Z-stack positions with the newly obtained XY position of the center of the sample.</li>
	<li>Turn both Z-stack and Tile Scan modes on. Set Z step (in Z-stack pane) to 5 &#181;m. Set Scanning Speed to 6. Turn the Autosave function on. Turn the option Saving Separate Tiles on. Name the file. Press the Start Experiment button.</li>
	<li>Verify that the experiment does not exceed the allocated time on the microscope; if so - adjust the speed. 
	&#8203;NOTE: The approximate file size is more than 10 Gb; be sure that there is enough space on the hard drive.</li>
</ol>
5. Spectral unmixing and stitching
<ol>
	<li>Use the software for the initial image processing. For spectral unmixing, select the Unmixing option. Select two regions corresponding to the streptavidin/airways and conidia to acquire the spectra from the image. Click Start Unmixing. 
	&#8203;NOTE: Alternatively, use the existing spectra for 488 and 594 nm fluorochromes.</li>
	<li>Open the file for the image processing and select the Processing tab. In the Methods section, select Geometric and Stitching. In the Parameter section, select the New Output and mark the Fuse Tiles option. Use the reference mode with the selected channel corresponding to the airway fluorescence. Apply the Stitching by clicking Apply.</li>
</ol>
6. Image processing: surface rendering
<ol>
	<li>Open the image as a Surpass 3D View. Create an airway surface using the option Surface for the channel that was used for the airway visualization. Choose the Smoothing parameter of 10 &#181;m. 
	NOTE: Choose the automatic threshold value as well.</li>
	<li>Visually inspect the surface. Choose the thresholds to reduce the outlying signal. Remove the surfaces of pleura and vessels.</li>
	<li>Create a mask for the airway surface using options Edit and Mask All. Select the airway channel and set Voxels outside the surface to 0.001.</li>
	<li>Save the file with the airway mask as a TIFF series to the folder. Save the file with the conidia channel as a TIFF series to the separate folder.</li>
</ol>
7. Image processing: mask correction
<ol>
	<li>Open the file with the airway mask in an open-source platform for biological-image analysis17 as an 8-bit image. Make the image binary by clicking Process | Binary | Make Binary. 
	NOTE: If necessary, correct the mask: remove the excessive surfaces using the Polygon selection and Delete tab. Alternatively, use the Flood Fill Tool.</li>
	<li>Draw the missing surfaces using several times Dilate (3D): click Plugins | Process | Dilate (3D). Fill the holes by clicking Process | Binary | Fill Holes. Use selection and ROI manager interpolation to fill the residual holes in the mask manually. Apply several Erode (3D) options (Plugins | Process | Erode (3D)) to resample the mask thickness. 
	NOTE: Create macros by using Plugins | Macros options for multiple Dilate (3D) and Erode (3D) repeats. 
	Ensure the number of Erode (3D) is equal to the number of Dilate (3D) applications.</li>
	<li>Save the mask in a new folder as a TIFF series.</li>
</ol>
8. Conidia quantitative analysis
<ol>
	<li>Open the app in the programming and numeric computing platform: https://www.mathworks.com/matlabcentral/fileexchange/84525-conidia_counter.</li>
	<li>Press the Add files button. Select the airway mask folder and the conidia folder.</li>
	<li>Set a custom threshold between 0 and 1. Press the OK button.</li>
	<li>Save the output table to the custom .xls file.</li>
	<li>Analyze the data with software for statistical analysis.</li>
</ol>

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The authors report no conflicts of interest in this work.

Whole-organ 3D imaging permits obtaining of the data without dissection of the specimen, which is of great importance for investigating the spatial aspects of the anatomical distribution of the pathogen in the organism. There are several techniques and modifications of tissue optical clearing that help to overcome the laser light scattering and allow whole-organ imaging15,16,18,19. One of the c...

On a daily basis, people inhale airborne pathogens, including spores of opportunistic fungi Aspergillus fumigatus (A. fumigatus conidia) that can penetrate the respiratory tract1. The respiratory tract of mammals is a system of airways of different generations that are characterized by the different structures of the airway walls2,3,4. Tracheobronchial walls consist of several cell types among which are ciliated cells that provide the mucociliary clearance5. In the alveoli, there are no ciliated cells and the penetrating alveolar space pathogens cannot be eliminated by the mucociliary clearance6. Moreover, each airway generation is a niche for multiple immune cell populations and subsets of these populations are unique for certain airway compartments. Thus, alveolar macrophages reside in the alveolar compartments, while both the trachea and conducting airways are lined with the intraepithelial dendritic cells7,8.
The approximate size of A. fumigatus conidia is 2-3.5 &#181;m9. Since the diameter of small airways in humans and even in mice exceeds 3.5 &#181;m, it was suggested that conidia can penetrate the alveolar space2,10,11. In fact, histological examination showed the fungal growth in the alveoli of the patients suffering from aspergillosis12. Conidia were also detected in the alveoli of infected mice using live imaging of the thick lung slices13. Simultaneously, conidia were detected in the luminal side of the bronchial epithelium of mice14.
Three-dimensional (3D) imaging of the optically cleared whole-mount mouse lungs permits morphometric analysis of the airways15. Particularly, the quantitative analysis of the visceral pleural nerve distribution was performed using optically cleared mouse lung specimens15. Recently, Amich et al.16 investigated the fungal growth after intranasal application of conidia to the immunocompromised mice using a light-sheet fluorescence microscopy of optically cleared mouse lung specimens. The precise location of the resting conidia in the airways at different time points after the infection is important for identifying the cell populations that can provide sufficient antifungal defense in certain phases of inflammation. However, due to the relatively small size, the spatio-temporal aspects of A. fumigatus conidia distribution in the airways are poorly characterized.
Here, we present an experimental setup for the quantitative analysis of A. fumigatus conidia distribution in the airways of infected mice. Using fluorescent confocal laser scanning microscopy (CLSM) of optically cleared lungs of mice that received an oropharyngeal application of the fluorescently labeled A. fumigatus conidia, we obtain 3D images and perform the image processing. Using 3D imaging of the whole-mount lung lobe, we have previously shown the distribution of A. fumigatus conidia in the conducting airway of mice 72 hours after conidia application8.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Alexa Fluor 594 NHS Ester</td>
			<td>ThermoFisher</td>
			<td>A20004</td>
			<td></td>
		</tr>
		<tr>
			<td>Aspergillus fumigatus conidia</td>
			<td>ATCC</td>
			<td>46645</td>
			<td>The strain AfS150, a ATCC 46645 derivative</td>
		</tr>
		<tr>
			<td>Benzyl alcohol</td>
			<td>Panreac</td>
			<td>141081.1611</td>
			<td>98.0-100 %</td>
		</tr>
		<tr>
			<td>Benzyl benzoate</td>
			<td>Acros</td>
			<td>AC10586-0010</td>
			<td>99+%</td>
		</tr>
		<tr>
			<td>C57Bl/6 mice</td>
			<td>Pushchino Animal Breeding Centre (Russia)</td>
			<td></td>
			<td>Male. 12 - 30 week old.</td>
		</tr>
		<tr>
			<td>Catheter</td>
			<td>Venisystems</td>
			<td>G715-A01</td>
			<td>18G</td>
		</tr>
		<tr>
			<td>Cell imaging coverglass-bottom chamber</td>
			<td>Eppendorf</td>
			<td>30742028</td>
			<td>4 or 8 well chamber with coverglass bottom</td>
		</tr>
		<tr>
			<td>Centrifuge</td>
			<td>Eppendorf</td>
			<td>5804R</td>
			<td>Any centrifuge provided 1000 g can be used</td>
		</tr>
		<tr>
			<td>Confocal laser scanning microscope</td>
			<td>ZEISS</td>
			<td>ZEISS LSM780</td>
			<td></td>
		</tr>
		<tr>
			<td>Dimethyl sulfoxide</td>
			<td>Sigma-Aldrich</td>
			<td>276855</td>
			<td>&#8805;99.9%</td>
		</tr>
		<tr>
			<td>FIJI image processing package</td>
			<td>FIJI</td>
			<td></td>
			<td>Free software</td>
		</tr>
		<tr>
			<td>Forcep</td>
			<td>B. Braun Aesculap</td>
			<td>BD557R</td>
			<td>Toothed</td>
		</tr>
		<tr>
			<td>Forcep</td>
			<td>B. Braun Aesculap</td>
			<td>BD321R</td>
			<td>Fine-tipped</td>
		</tr>
		<tr>
			<td>Forcep</td>
			<td>Bochem</td>
			<td>1727</td>
			<td>Smooth</td>
		</tr>
		<tr>
			<td>Glass bottle</td>
			<td>DURAN</td>
			<td>242101304</td>
			<td>With groung-in lid</td>
		</tr>
		<tr>
			<td>Graphic Editor Photoshop</td>
			<td>Adobe Inc</td>
			<td></td>
			<td>Adobe Photoshop CS</td>
		</tr>
		<tr>
			<td>GraphPad Software</td>
			<td>GraphPad</td>
			<td></td>
			<td>Prism 8</td>
		</tr>
		<tr>
			<td>Imaris Microscopy Imaging Software</td>
			<td>Oxford Instruments</td>
			<td></td>
			<td>Free trial is avalable https://imaris.oxinst.com/microscopy-imaging-software-free-trial</td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Karizoo</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>Panreac</td>
			<td>141638</td>
			<td></td>
		</tr>
		<tr>
			<td>Objective</td>
			<td>ZEISS</td>
			<td>420640-9800-000</td>
			<td>&#160;Plan-Apochromat, 10 &#215; (NA = 0.3)</td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Sigma-Aldrich</td>
			<td>158127</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>Paneco</td>
			<td>P060&#928;</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette</td>
			<td>ProLine</td>
			<td>722020</td>
			<td>5 to 50 &#956;L</td>
		</tr>
		<tr>
			<td>Powdered milk</td>
			<td>Roth</td>
			<td>T145.2</td>
			<td></td>
		</tr>
		<tr>
			<td>Sample mixer</td>
			<td>Dynal</td>
			<td>MXIC1</td>
			<td></td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>B. Braun</td>
			<td>BC257R</td>
			<td>Blunt</td>
		</tr>
		<tr>
			<td>Shaker</td>
			<td>Apexlab</td>
			<td>GS-20</td>
			<td>50-300 rpm</td>
		</tr>
		<tr>
			<td>Skalpel</td>
			<td>Bochem</td>
			<td>12646</td>
			<td></td>
		</tr>
		<tr>
			<td>Silk thread</td>
			<td>B. Braun</td>
			<td></td>
			<td>3 USP</td>
		</tr>
		<tr>
			<td>Streptavidin, Alexa Fluor 488 conjugate</td>
			<td>ThermoFisher</td>
			<td>S11223</td>
			<td></td>
		</tr>
		<tr>
			<td>Test tube</td>
			<td>SPL Lifesciences</td>
			<td>50050</td>
			<td>50 mL</td>
		</tr>
		<tr>
			<td>Tris (hydroxymethyl aminomethane)</td>
			<td>Helicon</td>
			<td>H-1702-0.5</td>
			<td>&#160;Mr 121.14; CAS Number: 77-86-1</td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Amresco</td>
			<td>Am-O694-0.1</td>
			<td></td>
		</tr>
		<tr>
			<td>ZEN microscope software</td>
			<td>ZEISS</td>
			<td>ZEN2012 SP5</td>
			<td>https://www.zeiss.com/microscopy/int/products/microscope-software/zen.html</td>
		</tr>
	</tbody>
</table>

confocal laser scanning microscopy-based quantitative analysis of aspergillus fumigatus conidia distribution in whole-mount optically cleared mouse lung

Aspergillus fumigatus conidia are airborne pathogens that can penetrate human airways. Immunocompetent people without allergies exhibit resistance and immunological tolerance, while in immunocompromised patients, conidia can colonize airways and cause severe invasive respiratory disorders. Various cells in different airway compartments are involved in the immune response that prevents fungal invasion; however, the spatio-temporal aspects of pathogen elimination are still not completely understood. Three-dimensional (3D) imaging of optically cleared whole-mount organs, particularly the lungs of experimental mice, permits detection of fluorescently labeled pathogens in the airways at different time points after infection. In the present study, we describe an experimental setup to perform a quantitative analysis of A. fumigatus conidia distribution in the airways. Using fluorescent confocal laser scanning microscopy (CLSM), we traced the location of fluorescently labeled&#160;conidia in the bronchial branches and the alveolar compartment 6 hours after oropharyngeal application to mice. The approach described here was previously used for detection of the precise pathogen location and identification of the pathogen-interacting cells at different phases of the immune response. The experimental setup can be used to estimate the kinetics of the pathogen elimination in different pathological conditions.

Following the protocol above, the 3D image showing the airways and A. fumigatus conidia in the lung lobe of a mouse was obtained (Figure 1A). Streptavidin (that was used for airway visualization) labeled bronchi and bronchioles15. Additionally, the large vessels, which are easily distinguishable from the airways by their morphology, and pleura are visualized in the airway channel (Figure 1A-C). The creation of th...

Watch this Scientific Journal Video about Confocal Laser Scanning Microscopy-Based Quantitative Analysis of Aspergillus fumigatus Conidia Distribution in Whole-Mount Optically Cleared Mouse Lung at Jo

Confocal Laser Scanning Microscopy-Based Quantitative Analysis of Aspergillus fumigatus Conidia Distribution in Whole-Mount Optically Cleared Mouse Lung

We describe the method for quantitative analysis of the distribution of Aspergillus fumigatus conidia (3 &#181;m in size) in the airways of mice. The method also can be used for the analysis of microparticles and nanoparticle agglomerate distribution in the airways in various pathological condition models.

Confocal Laser Scanning Microscopy-Based Quantitative Analysis of Aspergillus fumigatus Conidia Distribution in Whole-Mount Optically Cleared Mouse Lung

Aspergillus fumigatus conidia are airborne pathogens that can penetrate human airways. Immunocompetent people without allergies exhibit resistance and ...

confocal-laser-scanning-microscopy-based-quantitative-analysis

Research

JoVE Journal

Immunology and Infection

3.2K Views.  Moscow Institute of Physics and Technology. We describe the method for quantitative analysis of the distribution of Aspergillus fumigatus conidia (3 µm in size) in the airways of mice. The method also can be used for the analysis of microparticles and nanoparticle agglomerate distribution in the airways in various pathological condition models.

Video: Confocal Laser Scanning Microscopy-Based Quantitative Analysis of Aspergillus fumigatus Conidia Distribution in Whole-Mount Optically Cleared Mouse Lung

Isolation of Mouse Lung Dendritic Cells

Lung dendritic cells (DC) play a fundamental role in sensing invading pathogens 1,2 as well as in the control of tolerogenic responses 3 in the respiratory tract. At least three main subsets of lung dendritic cells have been described in mice: conventional DC (cDC) 4, plasmacytoid DC (pDC) 5 and the IFN-producing killer DC (IKDC) 6,7. The cDC subset is the most prominent DC subset in the lung 8. 
The common marker known to identify DC subsets is CD11c, a type I transmembrane integrin (&beta;2) that is also expressed on monocytes, macrophages, neutrophils and some B cells 9. In some tissues, using CD11c as a marker to identify mouse DC is valid, as in spleen, where most CD11c+ cells represent the cDC subset which expresses high levels of the major histocompatibility complex class II (MHC-II). However, the lung is a more heterogeneous tissue where beside DC subsets, there is a high percentage of a distinct cell population that expresses high levels of CD11c bout low levels of MHC-II. Based on its characterization and mostly on its expression of F4&#47;80, an splenic macrophage marker, the CD11chiMHC-IIlo lung cell population has been identified as pulmonary macrophages 10 and more recently, as a potential DC precursor 11. 
In contrast to mouse pDC, the study of the specific role of cDC in the pulmonary immune response has been limited due to the lack of a specific marker that could help in the isolation of these cells. Therefore, in this work, we describe a procedure to isolate highly purified mouse lung cDC. The isolation of pulmonary DC subsets represents a very useful tool to gain insights into the function of these cells in response to respiratory pathogens as well as environmental factors that can trigger the host immune response in the lung.

A highly purified preparation of mouse lung dendritic cells is described. Specific emphasis is given to the isolation of conventional dendritic cell subset.

Lung dendritic cells (DC) play a fundamental role in sensing invading pathogens 1,2 as well as in the control of tolerogenic responses 3 in the ...

Intravital Microscopy of the Spleen: Quantitative Analysis of Parasite Mobility and Blood Flow

The advent of intravital microscopy in experimental rodent malaria models has allowed major advances to the knowledge of parasite-host interactions 1,2. Thus, in vivo imaging of malaria parasites during pre-erythrocytic stages have revealed the active entrance of parasites into skin lymph nodes 3, the complete development of the parasite in the skin 4, and the formation of a hepatocyte-derived merosome to assure migration and release of merozoites into the blood stream 5. Moreover, the development of individual parasites in erythrocytes has been recently documented using 4D imaging and challenged our current view on protein export in malaria 6. Thus, intravital imaging has radically changed our view on key events in Plasmodium development. Unfortunately, studies of the dynamic passage of malaria parasites through the spleen, a major lymphoid organ exquisitely adapted to clear infected red blood cells are lacking due to technical constraints.
Using the murine model of malaria Plasmodium yoelii in Balb/c mice, we have implemented intravital imaging of the spleen and reported a differential remodeling of it and adherence of parasitized red blood cells (pRBCs) to barrier cells of fibroblastic origin in the red pulp during infection with the non-lethal parasite line P.yoelii 17X as opposed to infections with the P.yoelii 17XL lethal parasite line 7. To reach these conclusions, a specific methodology using ImageJ free software was developed to enable characterization of the fast three-dimensional movement of single-pRBCs. Results obtained with this protocol allow determining velocity, directionality and residence time of parasites in the spleen, all parameters addressing adherence in vivo. In addition, we report the methodology for blood flow quantification using intravital microscopy and the use of different colouring agents to gain insight into the complex microcirculatory structure of the spleen. 
Ethics statement
All the animal studies were performed at the animal facilities of University of Barcelona in accordance with guidelines and protocols approved by the Ethics Committee for Animal Experimentation of the University of Barcelona CEEA-UB (Protocol No DMAH: 5429). Female Balb/c mice of 6-8 weeks of age were obtained from Charles River Laboratories.

We show the method for performing intravital microscopy of the spleen using GFP transgenic malaria parasites and the quantification of parasite mobility and blood flow within this organ.

The advent of intravital microscopy in experimental rodent malaria models has allowed major advances to the knowledge of parasite-host interactions 1,2. ...

Legionella pneumophila Outer Membrane Vesicles: Isolation and Analysis of Their Pro-inflammatory Potential on Macrophages

Bacteria are able to secrete a variety of molecules via various secretory systems. Besides the secretion of molecules into the extracellular space or directly into another cell, Gram-negative bacteria can also form outer membrane vesicles (OMVs). These membrane vesicles can deliver their cargo over long distances, and the cargo is protected from degradation by proteases and nucleases. 
Legionella pneumophila (L. pneumophila) is an intracellular, Gram-negative pathogen that causes a severe form of pneumonia. In humans, it infects alveolar macrophages, where it blocks lysosomal degradation and forms a specialized replication vacuole. Moreover, L. pneumophila produces OMVs under various growth conditions. To understand the role of OMVs in the infection process of human macrophages, we set up a protocol to purify bacterial membrane vesicles from liquid culture. The method is based on differential ultracentrifugation. The enriched OMVs were subsequently analyzed with regard to their protein and lipopolysaccharide (LPS) amount and were then used for the treatment of a human monocytic cell line or murine bone marrow-derived macrophages. The pro-inflammatory responses of those cells were analyzed by enzyme-linked immunosorbent assay. Furthermore, alterations in a subsequent infection were analyzed. To this end, the bacterial replication of L. pneumophila in macrophages was studied by colony-forming unit assays. 
Here, we describe a detailed protocol for the purification of L. pneumophila OMVs from liquid culture by ultracentrifugation and for the downstream analysis of their pro-inflammatory potential on macrophages.

Legionella pneumophila Outer Membrane Vesicles: Isolation and Analysis of Their Pro-inflammatory Potential on Macrophages

Here, we describe the purification of Legionella pneumophila (L. pneumophila) outer membrane vesicles (OMVs) from liquid cultures. These purified vesicles are then used for the treatment of macrophages to analyze their pro-inflammatory potential.

Bacteria are able to secrete a variety of molecules via various secretory systems. Besides the secretion of molecules into the extracellular space or ...

Human Lung Dendritic Cells: Spatial Distribution and Phenotypic Identification in Endobronchial Biopsies Using Immunohistochemistry and Flow Cytometry

The lungs are constantly exposed to the external environment, which in addition to harmless particles, also contains pathogens, allergens, and toxins. In order to maintain tolerance or to induce an immune response, the immune system must appropriately handle inhaled antigens. Lung dendritic cells (DCs) are essential in maintaining a delicate balance to initiate immunity when required without causing collateral damage to the lungs due to an exaggerated inflammatory response. While there is a detailed understanding of the phenotype and function of immune cells such as DCs in human blood, the knowledge of these cells in less accessible tissues, such as the lungs, is much more limited, since studies of human lung tissue samples, especially from healthy individuals, are scarce. This work presents a strategy to generate detailed spatial and phenotypic characterization of lung tissue resident DCs in healthy humans that undergo a bronchoscopy for the sampling of endobronchial biopsies. Several small biopsies can be collected from each individual and can be subsequently embedded for ultrafine sectioning or enzymatically digested for advanced flow cytometric analysis. The outlined protocols have been optimized to yield maximum information from small tissue samples that, under steady-state conditions, contain only a low frequency of DCs. While the present work focuses on DCs, the methods described can directly be expanded to include other (immune) cells of interest found in mucosal lung tissue. Furthermore, the protocols are also directly applicable to samples obtained from patients suffering from pulmonary diseases where bronchoscopy is part of establishing the diagnosis, such as chronic obstructive pulmonary disease (COPD), sarcoidosis, or lung cancer.

Lung-resident immune cells, including dendritic cells (DCs) in humans, are critical for defense against inhaled pathogens and allergens. However, due to the scarcity of human lung tissue, studies are limited. This work presents protocols to process human mucosal endobronchial biopsies for studying lung DCs using immunohistochemistry and flow cytometry.

The lungs are constantly exposed to the external environment, which in addition to harmless particles, also contains pathogens, allergens, and toxins. In ...

Live Imaging of Antifungal Activity by Human Primary Neutrophils and Monocytes in Response to A. fumigatus

Aspergillus fumigatus is an opportunistic fungal pathogen causing invasive infections in immunocompromised hosts with a high case-fatality rate. Research investigating immunological responses against A. fumigatus has been limited by the lack of consistent and reliable assays for measuring the antifungal activity of specific immune cells in vitro. A new method is described to assess the antifungal activity of primary monocytes and neutrophils from human donors against A. fumigatus using FLuorescent Aspergillus REporter (FLARE) conidia. These conidia contain a genetically encoded dsRed reporter, which is constitutively expressed by live FLARE conidia, and are externally labeled with Alexa Fluor 633, which is resistant to degradation within the phagolysosome, thus allowing a distinction between live and dead A. fumigatus conidia. Video microscopy and flow cytometry are subsequently used to visualize the interaction between conidia and innate immune cells, assessing fungicidal activity whilst also providing a wealth of information on phagocyte migration, phagocytosis and the inhibition of fungal growth. This novel technique has already provided exciting new insights into the host-pathogen interaction of primary immune cells against A. fumigatus. It is important to note the laboratory setup required to perform this assay, including the necessary microscopy and flow cytometry facilities, and the capacity to work with human donor blood and genetically manipulated fungi. However, this assay is capable of generating large amounts of data and can reveal detailed insights into the antifungal response. This protocol has successfully been used to study the host-pathogen interaction of primary immune cells against A. fumigatus.
It is important to note the laboratory setup required to perform this assay, including the necessary microscopy and flow cytometry facilities, and the capacity to work with human donor blood and genetically manipulated fungi. However, this assay is capable of generating large amounts of data and can reveal detailed insights into the antifungal response. This protocol has successfully been used to study the host-pathogen interaction of primary immune cells against A. fumigatus.

Live Imaging of Antifungal Activity by Human Primary Neutrophils and Monocytes in Response to A. fumigatus

Here, we describe a protocol to assess antifungal activity of primary human immune cells in real-time using fluorescent Aspergillus reporter conidia in conjunction with live-cell video microscopy and flow cytometry. Generated data provide insight into host cell-Aspergillus interactions such as fungicidal activity, phagocytosis, cell migration and inhibition of fungal growth.

Aspergillus fumigatus is an opportunistic fungal pathogen causing invasive infections in immunocompromised hosts with a high case-fatality rate. Research ...

Optical Screening of Novel Bacteria-specific Probes on Ex Vivo Human Lung Tissue by Confocal Laser Endomicroscopy

Improving the speed and accuracy of bacterial detection is important for patient stratification and to ensure the appropriate use of antimicrobials. To achieve this goal, the development of diagnostic techniques to recognize bacterial presence in real-time at the point-of-care is required. Optical imaging for direct identification of bacteria within the host is an attractive approach. Several attempts at chemical probe design and validation have been investigated, however none have yet been successfully translated into the clinic. Here we describe a method for ex vivo validation of bacteria-specific probes for identification of bacteria within the distal lung, imaged by fibered confocal fluorescence microscopy (FCFM). Our model used ex vivo human lung tissue and a clinically approved confocal laser endomicroscopy (CLE) platform to screen novel bacteria-specific imaging compounds, closely mimicking imaging conditions expected to be encountered with patients. Therefore, screening compounds by this technique provides confidence of potential clinical tractability.

Optical Screening of Novel Bacteria-specific Probes on Ex Vivo Human Lung Tissue by Confocal Laser Endomicroscopy

This technique describes an efficient screening process for evaluating bacteria-specific optical imaging agents within ex vivo human lung tissue, by fibered confocal fluorescence microscopy for the rapid identification of small molecule chemical probe-candidates with translatable potential.

Improving the speed and accuracy of bacterial detection is important for patient stratification and to ensure the appropriate use of antimicrobials. To ...

Visualizing Macrophage Extracellular Traps Using Confocal Microscopy

A primary method used to define the presence of neutrophil extracellular traps (NETs) is confocal microscopy. We have modified established confocal microscopy methods to visualize macrophage extracellular traps (METs). These extracellular traps are defined by the presence of extracellular chromatin with co-expression of other components such as granule proteases, citrullinated histones, and peptidyl arginase deiminase (PAD). The expression of METs is generally measured after exposure to a stimulus and compared to un-stimulated samples. Samples are also included for background and isotype control. Cells are analyzed using well-defined image analysis software. Confocal microscopy may be used to define the presence of METs both in vitro and in vivo in lung tissue.

Macrophage extracellular traps are a newly described entity. This article will concentrate on confocal microscopy methods and how they are visualized in vitro and in vivo from lung samples.

A primary method used to define the presence of neutrophil extracellular traps (NETs) is confocal microscopy. We have modified established confocal ...

Assessment of the Cytotoxic and Immunomodulatory Effects of Substances in Human Precision-cut Lung Slices

Respiratory diseases in their broad diversity need appropriate model systems to understand the underlying mechanisms and enable development of new therapeutics. Additionally, registration of new substances requires appropriate risk assessment with adequate testing systems to avoid the risk of individuals being harmed, for example, in the working environment. Such risk assessments are usually conducted in animal studies. In view of the 3Rs principle and public skepticism against animal experiments, human alternative methods, such as precision-cut lung slices (PCLS), have been evolving. The present paper describes the ex vivo technique of human PCLS to study the immunomodulatory potential of low-molecular-weight substances, such as ammonium hexachloroplatinate (HClPt). Measured endpoints include viability and local respiratory inflammation, marked by altered secretion of cytokines and chemokines. Pro-inflammatory cytokines, tumor necrosis factor alpha (TNF-&#945;), and interleukin 1 alpha (IL-1&#945;) were significantly increased in human PCLS after exposure to a sub-toxic concentration of HClPt. Even though the technique of PCLS has been substantially optimized over the past decades, its applicability for the testing of immunomodulation is still in development. Therefore, the results presented here are preliminary, even though they show the potential of human PCLS as a valuable tool in respiratory research.

In view of the 3Rs principle, respiratory models as alternatives to animal studies are evolving. Especially for risk assessment of respiratory substances, there is a lack of appropriate assays. Here, we describe the use of human precision-cut lung slices for the assessment of airborne substances.

Respiratory diseases in their broad diversity need appropriate model systems to understand the underlying mechanisms and enable development of new ...

Induction of Mouse Lung Injury by Endotracheal Injection of Bleomycin

Pulmonary fibrosis is a hallmark of several human lung diseases with a different etiology. Since current therapies are rather limited, mouse models continue to be an essential tool for developing new antifibrotic strategies. Here we provide an effective method to investigate in vivo antifibrotic activity of human mesenchymal stromal cells obtained from whole umbilical cord (hUC-MSC) in attenuating bleomycin-induced lung injury. C57BL/6 mice receive a single endotracheal injection of bleomycin (1.5 U/kg body weight) followed by a double infusion of hUC-MSC (2.5 x 105) into the tail vein, 24 h and 7 days after the bleomycin administration. Upon sacrifice at days 8, 14, or 21, inflammatory and fibrotic changes, collagen content, and hUC-MSC presence in explanted lung tissue are analyzed. The injection of bleomycin into the mouse&#160;trachea allows the direct targeting of the lungs, leading to extensive pulmonary inflammation and fibrosis. The systemic administration of a double dose of hUC-MSC results in the early blunting of the bleomycin-induced lung injury. Intravenously infused hUC-MSC are transiently engrafted into the mouse lungs, where they exert their anti-inflammatory and antifibrotic activity. In conclusion, this protocol has been successfully applied for the preclinical testing of hUC-MSC in an experimental mouse model of human pulmonary fibrosis. However, this technique can be easily extended both to study the effect of different endotracheally administered substances on the pathophysiology of the lungs and to validate new anti-inflammatory and antifibrotic systemic therapies.

Here we present an effective method to investigate the antifibrotic activity of intravenously infused human mesenchymal stromal cells obtained from the whole umbilical cord following the induction of lung injury by an endotracheal injection of bleomycin in C57BL/6 mice. This protocol can be easily extended to the preclinical testing of other therapeutics.

Pulmonary fibrosis is a hallmark of several human lung diseases with a different etiology. Since current therapies are rather limited, mouse models ...

Measuring Phagocytosis of Aspergillus fumigatus Conidia by Human Leukocytes using Flow Cytometry

Invasive pulmonary infection by the mold Aspergillus fumigatus poses a great threat to immunocompromised patients. Inhaled fungal conidia (spores) are cleared from the human lung alveoli by being phagocytosed by innate monocytes and/or neutrophils. This protocol offers a fast and reliable measurement of phagocytosis by flow cytometry using fluorescein isothiocyanate (FITC)-labeled conidia for co-incubation with human leukocytes and subsequent counterstaining with an anti-FITC antibody to allow discrimination of internalized and cell-adherent conidia. Major advantages of this protocol are its rapidness, the possibility to combine the assay with cytometric analysis of other cell markers of interest, the simultaneous analysis of monocytes and neutrophils from a single sample and its applicability to other cell wall-bearing fungi or bacteria. Determination of percentages of phagocytosing leukocytes provides a means to microbiologists for evaluating virulence of a pathogen or for comparing pathogen wildtypes and mutants as well as to immunologists for investigating human leukocyte capabilities to combat pathogens.

Measuring Phagocytosis of Aspergillus fumigatus Conidia by Human Leukocytes using Flow Cytometry

This protocol provides a fast and reliable method to quantitatively measure phagocytosis of Aspergillus fumigatus conidia by human primary phagocytes using flow cytometry and to discriminate phagocytosis of conidia from mere adhesion to leukocytes.

Invasive pulmonary infection by the mold Aspergillus fumigatus poses a great threat to immunocompromised patients. Inhaled fungal conidia (spores) are ...