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

Summary

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.

Abstract

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 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.

Introduction

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 walls^2,3,4. 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 cells^7,8.

The approximate size of A. fumigatus conidia is 2-3.5 µm⁹. Since the diameter of small airways in humans and even in mice exceeds 3.5 µm, it was suggested that conidia can penetrate the alveolar space^2,10,11. 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.

Protocol

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

1. To obtain fluorescently labeled A. fumigatus conidia, fix 5 × 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.
2. 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 fresh PBS in a volume of 20 mL. Repeat.
3. Dissolve the conidia in 900 µL of 0.1 M NaHCO₃.
4. Dissolve the succinimidyl ester of the 594 nm fluorescent dye in 100 µL of dimethyl sulfoxide and add to conidia.
5. Incubate the conidia with the dye for 1 h on a shaker at 150 rpm at room temperature.
6. Wash the conidia twice with 20 mL of PBS in the centrifuge at 1,000 x g for 15 min.
7. Dilute the conidia to a concentration of 1 × 10⁸ conidia/mL in PBS and store at 4 °C.
8. 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 µL of conidia suspension to the mouse pharynx. Wait until the suspension is inhaled.

2. Specimen preparation

1. 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.
2. 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.
   1. Treat the mouse with 70% ethanol using a sprayer.
   2. Make a median longitudinal cut in the ventral skin from the hind paws to the forepaws and the chin.
   3. Separate the skin from the subcutaneous tissue using toothed forceps and closed scissors. Fix the upper ends of the skin with needles.
   4. 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.
   5. Make a medial cut of the thorax and neck connective tissues until the trachea is visible.
3. Run a silk thread beneath the trachea and make a surgical knot using two forceps.
   NOTE: Alternatively, use dental floss instead of silk thread.
   1. Carefully pull the thread and cut off the lungs from the connective tissue with scissors. Hold the scissors perpendicular to the table.
   2. 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 °C.
4. Dissect the lung lobes from the heart and each other with a scalpel.
   1. 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.
   2. 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.
   3. 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).
5. 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 °C for post-fixation.

3. Mouse lung lobe optical clearing

1. 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.
2. 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.
3. 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.
4. Put the specimen in the cell imaging coverglass-bottom chamber. The sample is ready for imaging.

4. Mouse lung lobe imaging with CLSM

1. 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.
2. 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.
3. Switch the software to the Acquisition tab. Select CLSM λ-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 µW). Adjust the Detector Gain between 750-900 range.
   NOTE: Higher gain settings are undesirable due to noise.
4. Narrow the pinhole to 1 Airy unit by clicking 1 AU button. Set pixel resolution to 512 × 512 pixels.
5. 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.
6. 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.
7. Position the objective using the focusing wheel near the bottom of the specimen, by looking at the Z-stack pane. This is approximately 20 µm from the bottom of the specimen.
8. 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 × 5 tiles are a good starting point.
9. 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.
10. Turn both Z-stack and Tile Scan modes on. Set Z step (in Z-stack pane) to 5 µm. 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.
11. Verify that the experiment does not exceed the allocated time on the microscope; if so - adjust the speed.
    ​NOTE: The approximate file size is more than 10 Gb; be sure that there is enough space on the hard drive.

5. Spectral unmixing and stitching

1. 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.
   ​NOTE: Alternatively, use the existing spectra for 488 and 594 nm fluorochromes.
2. 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.

6. Image processing: surface rendering

1. 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 µm.
   NOTE: Choose the automatic threshold value as well.
2. Visually inspect the surface. Choose the thresholds to reduce the outlying signal. Remove the surfaces of pleura and vessels.
3. 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.
4. 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.

7. Image processing: mask correction

1. 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.
2. 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.
3. Save the mask in a new folder as a TIFF series.

8. Conidia quantitative analysis

1. Open the app in the programming and numeric computing platform: https://www.mathworks.com/matlabcentral/fileexchange/84525-conidia_counter.
2. Press the Add files button. Select the airway mask folder and the conidia folder.
3. Set a custom threshold between 0 and 1. Press the OK button.
4. Save the output table to the custom .xls file.
5. Analyze the data with software for statistical analysis.

Results

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...

Discussion

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 imaging^15,16,18,19. One of the c...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Alexa Fluor 594 NHS Ester	ThermoFisher	A20004
Aspergillus fumigatus conidia	ATCC	46645	The strain AfS150, a ATCC 46645 derivative
Benzyl alcohol	Panreac	141081.1611	98.0-100 %
Benzyl benzoate	Acros	AC10586-0010	99+%
C57Bl/6 mice	Pushchino Animal Breeding Centre (Russia)		Male. 12 - 30 week old.
Catheter	Venisystems	G715-A01	18G
Cell imaging coverglass-bottom chamber	Eppendorf	30742028	4 or 8 well chamber with coverglass bottom
Centrifuge	Eppendorf	5804R	Any centrifuge provided 1000 g can be used
Confocal laser scanning microscope	ZEISS	ZEISS LSM780
Dimethyl sulfoxide	Sigma-Aldrich	276855	≥99.9%
FIJI image processing package	FIJI		Free software
Forcep	B. Braun Aesculap	BD557R	Toothed
Forcep	B. Braun Aesculap	BD321R	Fine-tipped
Forcep	Bochem	1727	Smooth
Glass bottle	DURAN	242101304	With ground-in lid
Graphic Editor Photoshop	Adobe Inc		Adobe Photoshop CS
GraphPad Software	GraphPad		Prism 8
Imaris Microscopy Imaging Software	Oxford Instruments		Free trial is available https://imaris.oxinst.com/microscopy-imaging-software-free-trial
Isoflurane	Karizoo
NaHCO3	Panreac	141638
Objective	ZEISS	420640-9800-000	Plan-Apochromat, 10 × (NA = 0.3)
Paraformaldehyde	Sigma-Aldrich	158127
PBS	Paneco	P060Π
Pipette	ProLine	722020	5 to 50 μL
Powdered milk	Roth	T145.2
Sample mixer	Dynal	MXIC1
Scissors	B. Braun	BC257R	Blunt
Shaker	Apexlab	GS-20	50-300 rpm
Skalpel	Bochem	12646
Silk thread	B. Braun		3 USP
Streptavidin, Alexa Fluor 488 conjugate	ThermoFisher	S11223
Test tube	SPL Lifesciences	50050	50 mL
Tris (hydroxymethyl aminomethane)	Helicon	H-1702-0.5	Mr 121.14; CAS Number: 77-86-1
Triton X-100	Amresco	Am-O694-0.1
ZEN microscope software	ZEISS	ZEN2012 SP5	https://www.zeiss.com/microscopy/int/products/microscope-software/zen.html