Name: in vivo evaluation of mucociliary clearance in mice
Uploaded: 2020-12-18T15:00:00.000+00:00
Duration: 6 min 35 s
Description: Watch this Scientific Journal Video about In vivo Evaluation of Mucociliary Clearance in Mice at JoVE.com

M.Z. and K.S.F. and this work was supported by a grant awarded under the Pitt Innovation Challenge (PInCh), through the Clinical and Translational Science Institute at the University of Pittsburgh, and NHLBI R01 grant HL153407, awarded to M.Z.

The University of Pittsburgh&#8217;s Institutional Animal Care and Use Committee approved all animal protocols specified in this publication prior to undertaking any of these animal experiments.
NOTE: This protocol details how to perform in vivo mucociliary clearance studies utilizing radionuclide imaging with a dual-modality SPECT/CT scanner. The techniques demonstrated are running system calibrations, anesthetizing mice, tracheal intubation of mice, instilling the isotope into the lungs, dual-modality imaging, co-registration of these images, and analysis.
1. SPECT/CT system setup
<ol>
	<li>Design an appropriate workflow and set up prior to running experiments using living animals.
	<ol>
		<li>Use a SPECT acquisition consisting of 60 projections with a step size of 6o between projections with a 40 cm radius of rotation. The CT acquisition consists of 220 projections with a 1.6o angle between projections.</li>
	</ol>
	</li>
	<li>Ensure that the system has the correct MWB collimators for mice and SPECT imaging in place. If the inappropriate collimators are installed, use the collimator wizard to install the correct ones.</li>
	<li>Run the necessary system calibrations to prepare the system for use. 
	NOTE: The SPECT and CT components of the scanner need calibration. Calibrate the CT components using a source conditioning and a Dark/Light (D/L) calibration once a day, a Center Offset (COS) calibration every 2 weeks, and evaluate the x-ray hardware every month. The SPECT components need to be calibrated once a year.
	<ol>
		<li>To evaluate the x-ray hardware, check the Evaluate X-ray hardware box during CT calibrations (Supplemental CT Calibration Menu).</li>
		<li>To perform source conditioning, check the Perform source conditioning box during CT calibrations (Supplemental CT Calibration Menu).</li>
		<li>To perform a D/L calibration, check the D/L box next to the CT acquisition protocol used during experiments during CT calibrations. Uncheck all other protocols (Supplemental CT Calibration Menu).</li>
		<li>To perform a COS calibration, replace the bed with the calibration ring tool, adjust the bed type settings to match in the motion control settings, and check the COS box next to the CT acquisition protocol used during experiments during CT calibrations. Uncheck all other protocols (Supplemental CT Calibration Menu, Supplemental Calibration Ring).</li>
	</ol>
	</li>
</ol>
2. Mouse Intubation and Instillation
<ol>
	<li>Weigh the mice to be scanned. If scanning multiple mice, take care to mark the mice for identification purposes using methods such as ear punching or marking of the tail.</li>
	<li>Anesthetize a mouse using 1.5% isoflurane with a gas flow of 2 L/min O2 in a gas chamber for ~5 minutes to produce anesthesia of sufficient depth, until breathing slows to ~55-65 breaths per minute 16 (Figure 1A).</li>
	<li>Remove the mouse from the chamber and suspend by the front incisors on an intubation stand at a 45o incline. Equip the intubation stand with a nose cone to ensure the mouse is anesthetized during the intubation (Figure 1B).</li>
	<li>Connect one end of a 50 &#181;m fiber optic wire to a light source and thread a 20-gauge cannula over it using the wire to act as a guide (Figure 1C).</li>
	<li>Open the mouth of the mouse and pull the tongue forward using blunt forceps. Illuminate the guide wire and use it to visualize the vocal cords (Figure 1D).</li>
	<li>Pass the guide wire through the vocal cords so that the wire is just beyond the vocal cords and resting in the upper trachea. Slide the 1 inch cannula forward along the wire to intubate the mouse, passing the cannula deep enough so that the hub of it is against the animal&#8217;s incisors (Figure 1E). Remove the wire leaving the cannula in place.</li>
	<li>Test the intubation by briefly plugging the cannula with a finger and checking for changes in breathing. Halted breathing or strained breathing while plugging and accelerated breathing upon release are signs of proper tracheal intubation. If there is no change in respiratory patterns upon plugging the cannula, the latter is likely in the esophagus.</li>
	<li>Prepare 0.2 mCi of 99mtechnetium-sulfur colloid (99mTc-Sc) in a volume of 10 &#181;L, and pipette into the cannula. Allow the mouse to spontaneously inhale it into the lungs over 1-2 min (Figure 1F). Remove the cannula before transferring the mouse to the pallet of the scanner. 
	NOTE: The radionuclide was prepared and filtered by Cardinal Health.</li>
</ol>
3. SPECT/CT Imaging
<ol>
	<li>Transfer the mouse to a 25 mm pallet with a nose cone and secure with tape, taking care to not tape the chest and abdomen too tightly to avoid impairing respiration. Take care to remove any metal ear tags attached to the mouse.</li>
	<li>Prepare a radioactive phantom consisting of 0.05 mCi in 200 &#181;L and place this amount in a 0.2 mL PCR tube. Position the tube by taping to the pallet under the lower abdomen of the mouse, avoiding overlap with the lungs. 
	NOTE: The phantom is used for the purpose of co-registering CT and SPECT images, as well as a negative control for clearance.</li>
	<li>Insert the mouse into the SPECT/CT system, select the imaging workflow, and run Setup.</li>
	<li>Set up the positioning of the detectors on the mouse, and run the imaging workflow.</li>
	<li>Prepare a cage for mice that have received radioactivity post-procedure, with unrestricted access to food and water, and clear labeling using a radiation safety sticker.</li>
	<li>Upon completion of the workflow, remove the mouse from the imaging pallet, and allow it to recover in the prepared cage for a duration of 6 hours between scans (end of scan 1 to beginning of scan 2) with ad libitum access to food and water. 6 h was chosen as it corresponds to the time period in which linear clearance depending on cilia function is taking place with very little alveolar clearance.</li>
	<li>After 6 hours, re-anaesthetize the mouse and scan, along with the phantom, using the same workflow to measure the amount of isotope cleared from the airways. 
	NOTE: It is critical to allow the mouse to recover as uninterrupted anesthesia with isoflurane for 6 hours will lead to a significant cilia-depressant effect, resulting in near zero mucociliary clearances.</li>
</ol>
4. Analysis
<ol>
	<li>After imaging, perform post-processing to reconstruct complete 3D stack images.
	<ol>
		<li>Histogram the SPECT images using the factory standard settings for 99mTc, and then reconstruct using a MAP3D algorithm and point spread function (PSF) reconstruction. 
		NOTE: The reconstruction was performed using 8 iterations and 6 subsets. An effective reconstruction needs a ratio of subsets to projections at 1:10 or divide evenly into the number of projections, so 6 subsets were used due to the acquisition using 60 projections.</li>
		<li>Reconstruct the CT images using the Feldkamp algorithm and a Shepp-Logan filter. 
		NOTE: The reconstruction was performed using 4 iterations.</li>
	</ol>
	</li>
	<li>Process the CT and SPECT images in FIJI ImageJ21 using the reslice tool to generate coronal view images from the default axial images. Then perform a z-stack sum projection on the SPECT image to add the count data from each slice and generate a single image for ease of analysis.</li>
	<li>Resize and co-register the CT and SPECT images using the phantom Eppendorf tube as a reference (Figure 2A,B). Track and use consistent resize measurements across all samples.</li>
	<li>Binarize the CT image using auto thresholding, followed by inverting the stack, and performing a z-stack sum projection to generate an outline of the lungs for analysis (Figure 2C).</li>
	<li>Rotate the CT and SPECT images and merge the image using the channel tools. Calculate MCC by drawing an ROI around the right lung and measuring (Figure 2D). 
	NOTE: This measurement will be of the total counts in the right lung for the 0 and 6 hour time points, with the 6 hour images corrected for radioactive decay using the formula: N(t) = N0e&#8722;t. 99mTc-Sc has a decay constant of 3.21e&#8722;5 per second with a half-life of ~6 hours. These values can then be used to calculate a percent clearance. 
	NOTE: Right lung is chosen for ROI drawing and measuring counts as the mucociliary clearance will transport the radioisotope out of the lungs to the pharynx from where it will be swallowed and end up in the stomach. Quite frequently, counts can be seen in the stomach that may overlap with the left lung and hence produce erroneous counts. This confounding can be avoided by measuring counts in the right lung only.</li>
</ol>

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None related to this work.

The role of motile respiratory cilia in both disease and development continues to evolve and be better appreciated. Synchronous, metachronal beating of multiple motile cilia on the apical surface of cells lining the tracheobronchial tree generate cephalad flow producing mucociliary clearance or MCC. MCC is compromised in ciliopathies like PCD22, acquired diseases like COPD18, and its importance is being recognized in CHDs, not traditionally considered to be ciliopathies. Recent data has shown respiratory ciliary dysfunction in both CHD with heterotaxy23 and without heterotaxy7. Such motile cilia dysfunction was shown to translate into greater respiratory symptoms9 as well as greater post-operative morbidity8. Most, if not all, of these diseases, have mouse models available and our protocol for measuring MCC in mice is a valuable tool that can be utilized to test potential therapeutics.
Animal models provide utility for understanding diseases and the development of therapies. In vivo animal imaging provides further utility with the ability to acquire multiple data points from the same animals, without the need to sacrifice the animals, allowing investigators to follow longitudinal course of disease as well as study duration of treatment effects. The mouse model of MCC has been developed over the course of decades by multiple investigators, initially being performed on beagle dogs using planar scintigraphy, a two dimensional nuclear imaging technique24. The technique was adapted for use in mice a decade later, followed by adaptation to SPECT imaging a decade after that25,26. The development of this technique in mouse models was a major development in the relevance of this technique, due to the availability of multiple mouse models of human diseases like PCD in which ciliary function is significantly altered. MCC has been assessed in mouse models of lung denervation and immunosuppression, and has the potential to be used in conjunction with other models19,26. MCC measurement studies in human patients with airway diseases such as CF, asthma, PCD, and ciliopathies associated with CHD have been conducted, and have yielded results that the technique can aid both studies of lung physiology and therapeutic efficacy13.
An important part of this protocol is setting up acquisitions with the correct imaging parameters to acquire accurate images for quantification. A number of factors are key when designing SPECT acquisition settings, including which collimators are used, the number of projections to acquire per revolution, and rotation step size. Collimator selection is a major factor in the sensitivity and resolution of the acquisition, and acquisition settings may need to be tailored to the collimator being used27. Alternately, when using bigger animals like rats, the collimators would need to be adjusted. Multiple pinhole collimators for example are more sensitive, but care should be taken when selecting a step size in order to avoid overlapping projections and causing undesired multiplexing, which can further increase sensitivity of the acquisition at the expense of some image ambiguity that can cause reconstruction artifacts25. Reconstruction setup is also key to generate quantifiable images. MAP3D is a commonly used iterative reconstruction algorithm, and PSF is a common reconstruction model. Both are reliable for reconstructing images, but care should be taken when setting the number of iterations and subsets. A higher number of iterations will increase the computational time required for the reconstruction, and increase the quality of the reconstruction with diminishing returns upon further increase.
In order to quantify images in ImageJ, the ideal measurement tool to use is RawIntDen, which outputs the sum value of pixels in a selection. When quantifying SPECT data across differently sized lung ROIs, the use of RawIntDen provides an absolute measure of counts and avoids adjusting the measurement to the area of the ROI, like the mean measurement would21.
This technique has a number of associated sources of error that the investigator should be cognizant of when applying this technique. A notable confounder is the use of anesthetic agents. Isoflurane is a fast acting, inhaled anesthetic that the mice recover from rapidly after completion of an acquisition. However, care should be taken to provide the mice with ample time to recover in their cages, and not kept anesthetized any longer than necessary. In our personal experience (unpublished data) mice that were kept anesthetized continuously using inhaled isoflurane between the 0 and 6 hour time-point showed negligible clearance. Likewise, a controlled dose of anesthetic is also necessary to ensure rapid recovery. When securing the animal to the pallet for imaging, the phantom tube used for co-registration should be kept low on the stomach to avoid artifacts from overlapping with the lungs. Likewise, to ensure a quality CT image, take care to remove any metal tags from the mouse to avoid artifacts from x-ray scattering.
The current MCC protocol can be applied to myriad animal models. This technique has a negligible effect on the health of the animal scanned, is well tolerated by mice, and because of this it can be used with disease models without risking the health of already delicate mice. The strength of this methodology comes from it being an in vivo technique, which allows for the acquisition of consistent and repeatable measurements of airway function without the sacrifice of animals to excise tracheas for video-microscopy, that ex vivo models require26. The consistency of this technique in producing repeatable measurements across multiple scans of the same animals, allows for the same animal to be treated with different agents or potential therapeutics, and statistical comparisons made between the same animal to reduce biological variability inherent in any animal model, thereby reducing the sample size needed to show statistically significant differences.
The assessment of airway function using the MCC technique can be adjusted to a variety of animal models and applied to many different models of airway health, as well as testing new therapies. The airways of mouse models of PCD can be assessed using this technique, as well as models of COPD. Our method can also be utilized to study differential effects of various anesthetics on MCC that are in common clinical use. Finally, the effects of therapeutic agents on the airway can also be assessed using this model. As previously stated but bears repetition, as it is an in vivo measurement it allows for repeat MCC assessments over the course of a disease, as well as test benefits of therapeutic interventions over time. Additionally, mice are the most common laboratory animals used to mimic/study human diseases, with, in some cases, multiple transgenic mouse models of human disease available to choose from.

Cilia are microtubule based cellular organelles conserved across evolutionary history from algae to humans. They emanate from cell surfaces and have a number of functions1, ranging from recognition of local environmental sensory signals to motility, functions which can be traced back from humans to early unicellular eukaryotic organisms2,3. Cilia can be non-motile and single serving as a cell&#8217;s specialized antenna to process environmental signals; or motile and multiple, beating in synchronized, metachronal waves to generate fluid flow, such as in the lining of the fallopian tubes and the upper and lower airways, except for the terminal bronchioles leading to the alveoli1,2.
The extensive epithelial surface of the respiratory tract is exposed to a constant barrage of contamination in the form of a variety of potentially hazardous inhaled pollutants and pathogens, necessitating a defense. One key defense mechanism is the mucociliary apparatus of the tracheobronchial tree, where a continuous flow of secreted mucus is mechanically transported out of the airway by the beating of multiple motile cilia lining the apical surfaces of the tracheo-bronchial epithelial cells. These function to entrap inhaled contaminants, and through their continuous, synchronal beating, transport them cephalad4,5.
Cilia have been demonstrated to have key roles such as in the development of left-right patterning in developing embryos, where motile cilia at the embryonic node break symmetry6. Mutations in cilia related genes have been linked to diseases such as congenital heart disease (CHD) due to the asymmetric structure of the heart6. Recent studies have reported a high incidence of ciliary dysfunction in the respiratory tracts of patients with CHD, as well as an increased prevalence of post-operative respiratory complications and chronic respiratory tract symptoms in the upper and lower airways7,8,9,10. Patients with CHD and ciliary dysfunction, with or without heterotaxy, have been demonstrated to have increased risk of respiratory complications and negative respiratory outcomes post-operatively5,8,10. Beyond their roles in signaling and development, the importance of airway cilia has been demonstrated by ciliopathies, of which a prime example is primary ciliary dyskinesia (PCD). PCD is a congenital disorder resulting from a number of mutations affecting the motile respiratory cilia, leading to recurrent lung infections, bronchiectasis, and potentially the need for lung transplantation11. Additionally, even though cilia are normal in cystic fibrosis (CF), most common congenital disorder in the Caucasian population, MCC is impaired due to thick, viscous mucus resulting from mutations in the CFTR gene12. There are multiple mouse models of PCD and CF, as well as an ever-increasing number of models of CHD. Ultimately cilia are versatile structures with many key roles, and a method to assess the function of motile respiratory cilia in vivo can be valuable for pre-clinical study, and assessing effects of mutations as well as drugs on mucociliary clearance (MCC)13. The method would also be valuable in assessing effects of novel drugs, gene therapy or interventions on MCC in these mouse models.
There are many different models that have been used to assess MCC. One notable method involves the use of methylene blue dye that has been instilled into the bronchus, with clearance measured by fiberoptic measurement of dye movement14. This method is limited by the ability to observe the movement of the dye, which is more routine in humans than in pre-clinical mouse models. Another notable method is synchrotron phase-contrast X-ray imaging (PCXI), which can be used to track individual particles in an airway. This method is relatively new and not widely accessible15. There are numerous ex vivo methods of assessing the airway by excising a trachea for video-microscopy, however these models provide little utility in human patients16. High resolution techniques for cilia imaging such as optical coherence tomography are limited in the same way17.
In this article, we present a reproducible method to measure MCC in vivo that has been used to measure lung clearances in myriad animal models, as well as study MCC in chronic obstructive pulmonary disease and assess the effects of immunosuppressive drugs18,19. This method tracks the clearance of the radiopharmaceutical 99mtechnetium-sulfur colloid (99mTc-Sc), an insoluble particulate radiotracer, after instillation into the lungs. The radionuclide can then be tracked using single photon emission computed tomography (SPECT)18,20. We have further refined this technique for measuring MCC by using dual modality SPECT and computed tomography (CT) imaging with co-localization of radioisotope counts to the lungs and measuring the decrease in these counts over 6 hours. Dual-modality imaging, with co-registration of CT and SPECT images allows for accurate localization of radiation counts to our region of interest, the lungs. Although we describe in detail the method for MCC measurement in mice, the protocol can be adjusted to study MCC in rats. The collimators would need to be adjusted as well as radiation dose. In our view, mouse MCC scans are more technically challenging due to the small animal size, but more useful than rats due to the large number of established mouse models of a number of human disorders. Additionally, due to their lower cost and cost of maintenance in animal colonies, a larger sample size is more feasible in mice.

<table><tbody><tr><td>500 &amp;micro;m Unjacketed Fiber Optic Wire</td><td>Edmund Optics</td><td>02-532</td><td></td></tr><tr><td>99mTechnecium-Sulfur Colloid</td><td>Cardinal Health</td><td></td><td></td></tr><tr><td>Anesthesia Vaporizer</td><td>Vetland Medical</td><td>A13480</td><td></td></tr><tr><td>Durmont #5 Forceps</td><td>Fine Science Tools</td><td>99150-20</td><td></td></tr><tr><td>FIJI ImageJ 2.0.0-rc-65/1.52p Software</td><td></td><td></td><td></td></tr><tr><td>Introcan Safety Catheters 20G 1inch</td><td>Fisher Scientific</td><td>NC1534477</td><td></td></tr><tr><td>Isoflurane</td><td>Henry Schein</td><td>118-2097</td><td></td></tr><tr><td>Mouse Intubation Stand</td><td>Kent Scientific</td><td>ETI-MSE-01</td><td></td></tr><tr><td>Siemens Inveon dual-modality SPECT/CT</td><td>Siemens</td><td></td><td></td></tr><tr><td>Single Channel Anesthesia Stand</td><td>Summit Anesthesia Solutions</td><td>22860</td><td></td></tr></tbody></table>

in vivo evaluation of mucociliary clearance in mice

Respiratory motile cilia, specialized organelles of the cell, line the apical surface of epithelial cells lining the respiratory tract. By beating in a metachronal, synchronal fashion, these multiple, motile, actin-based organelles generate a cephalad fluid flow clearing the respiratory tract of inhaled pollutants and pathogens. With increasing environmental pollution, novel viral pathogens and emerging multi-drug resistant bacteria, cilia generated mucociliary clearance (MCC) is essential for maintaining lung health. MCC is also depressed in multiple congenital disorders like primary ciliary dyskinesia, cystic fibrosis as well as acquired disorders like chronic obstructive pulmonary disease. All these disorders have established, in some case multiple, mouse models. In this publication, we detail a method using a small amount of radioactivity and dual-modality SPECT/CT imaging to accurately and reproducibly measure MCC in mice in vivo. The method allows for recovery of mice after imaging, making serial measurements possible, and testing potential therapeutics longitudinally over time. The data in wild-type mice demonstrates the reproducibility of the MCC measurement as long as adequate attention to detail is paid, and the protocol strictly adhered to.

Using this protocol, we anesthetized mice in an isoflurane chamber (Figure 1A). After achieving an adequate level of anesthesia, mice were placed onto vertical supports (Figure 1B) and the vocal cords were visualized using an illuminated guide wire (Figure 1C-1D). The mice were intubated and instilled with 0.2 mCi 99mTc-Sc in volumes of 10 &#181;L through a cannula and mice allowed to spontaneously inhale into the lungs (Figure 1E-1F). After image acquisition and processing, the CT and SPECT images were colocalized (Figure 2A) using the phantom tube as a landmark (Figure 2B). Masks of the lungs were generated from the CT image (Figure 2C) and used to draw ROIs around the right lung for analysis at 0 (Figure 2D) and 6 hours (Figure 2E-2F). To test reproducibility of the protocol, a total of 8 mice were scanned twice on different days with identical experimental conditions, with analysis using a paired t-test showing no significant difference between the repeat scans (p-value=0.9904) (Figure 3A). An additional 2 mice were scanned three times on different days with identical experimental conditions, with analysis using one-way ANOVA showing significant matching between the repeat scans (p-value of 0.0041) (Figure 3B). A total of 8 mice were scanned and two representative images were displayed (Figure 4).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61929/61929fig01.jpg" /> 
Figure 1: Mouse intubation and isotope instillation.&#160;Images of the steps needed to intubate and instill isotope into the airway. A) The mouse is anesthetized in a chamber. B) The anesthetized mouse is placed onto a vertical support, suspended by the front incisors. C) An illuminated 0.5 mm fiber optic wire serving as a guide wire is prepared by running it through a 20 G cannula. D) The mouth of the mouse is opened using forceps and illuminated using the lighted guidewire to visualize the vocal cords. E) The cannula is pushed across the vocal cords and guidewire is removed. F) Soluble isotope is instilled into the cannula using a pipette and the mouse allowed to spontaneously inhale the isotope into the lungs. <a href="https://www.jove.com/files/ftp_upload/61929/61929fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61929/61929fig02v2.jpg" /> 
Figure 2: SPECT/CT images of an MCC scan.&#160;A) A SPECT image that has been co-localized with a CT image. B) A CT image with a visible phantom tube that was used for co-localization. C) A mask of the airway derived by binarizing the CT image and performing a z-stack sum projection. D) The CT mask co-localized with the SPECT image. An ROI for analysis has been drawn around the right lung. E) A mask of the airway at 6 hours. F) A CT and SPECT co-localized image of the airway at 6 hours with an ROI for analysis. <a href="https://www.jove.com/files/ftp_upload/61929/61929fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61929/61929fig03.jpg" /> 
Figure 3: Clearance measurements of the same mice across multiple scans.&#160;A) Two individual repeat clearance were measured for 8 mice with no changes in experimental conditions. A paired t test showed that there was no significant difference between the repeat scans with a p-value of 0.9904. B) Three individual repeat clearances were measured for two mice with no changes in experimental conditions. A one-way ANOVA showed there was significant matching between the repeat scans with a p-value of 0.0041. <a href="https://www.jove.com/files/ftp_upload/61929/61929fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61929/61929fig04.jpg" /> 
Figure 4: Co-localized SPECT/CT images of the 0 and 6 hour airway in 2 mice with ROIs drawn at 0 and 6 hours outlining the right lung. <a href="https://www.jove.com/files/ftp_upload/61929/61929fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental Figure 1: A video of the vocal chords illuminated by a fiber optic wire with the effect of breathing visualized. <a href="https://cloudflare.jove.com/files/ftp_upload/61929/61929supfig01.jpg" target="_blank">Please click here to download this figure.</a>
Supplemental Files. <a href="https://cloudflare.jove.com/files/ftp_upload/61929/Supplemental_Files.zip" target="_blank">Please click here to download these files.</a>

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In vivo Evaluation of Mucociliary Clearance in Mice

In this publication, we describe protocols for assessing airway mucociliary clearance (MCC) in mice in vivo utilizing dual-modality radionuclide imaging. This protocol is designed for a single photon emission computed tomography (SPECT) and computed tomography (CT) acquisition protocol using mouse whole body (MWB) collimators in a dual SPECT/CT system.

Respiratory motile cilia, specialized organelles of the cell, line the apical surface of epithelial cells lining the respiratory tract. By beating in a ...

in-vivo-evaluation-of-mucociliary-clearance-in-mice

Nanyang Technological University

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4.3K Views.  University of Pittsburgh School of Medicine. In this publication, we describe protocols for assessing airway mucociliary clearance (MCC) in mice in vivo utilizing dual-modality radionuclide imaging. This protocol is designed for a single photon emission computed tomography (SPECT) and computed tomography (CT) acquisition protocol using mouse whole body (MWB) collimators in a dual SPECT/CT system.

Video: In vivo Evaluation of Mucociliary Clearance in Mice

Murine Model of Allergen Induced Asthma

Asthma is a major cause of morbidity and mortality, affecting some 300 million people throughout the world.1 More than 8% of the US population has asthma, with the prevalence increasing.2 As with other diseases, animal models of allergic airway disease greatly facilitate understanding of the underlying pathophysiology, help identify potential therapeutic targets, and allow preclinical testing of possible new therapies. Models of allergic airway disease have been developed in several animal species, but murine models are particularly attractive due to the low cost, ready availability, and well-characterized immune systems of these animals.3 Availability of a variety of transgenic strains further increases the attractiveness of these models.4 Here we describe two murine models of allergic airway disease, both employing ovalbumin as the antigen. Following initial sensitization by intraperitoneal injection, one model delivers the antigen challenge by nebulization, the other by intratracheal delivery. These two models offer complementary advantages, with each mimicking the major features of human asthma.5
The major features of acute asthma include an exaggerated airway response to stimuli such as methacholine (airway hyperresponsiveness; AHR) and eosinophil-rich airway inflammation. These are also prominent effects of allergen challenge in our murine models,5,6 and we describe techniques for measuring them and thus evaluating the effects of experimental manipulation. Specifically, we describe both invasive7 and non-invasive8 techniques for measuring airway hyperresponsiveness as well as methods for assessing infiltration of inflammatory cells into the airways and the lung. Airway inflammatory cells are collected by bronchoalveolar lavage while lung histopathology is used to assess markers of inflammation throughout the organ. These techniques provide powerful tools for studying asthma in ways that would not be possible in humans.

Experimental mouse models of allergic asthma offer new possibilities for studying disease pathogenesis and developing new therapeutics. These models are well suited to measuring factors governing the allergic immune response, airway inflammation, and pulmonary pathophysiology.

Asthma is a major cause of morbidity and mortality, affecting some 300 million people throughout the world.1 More than 8% of the US population has asthma, ...

Immunology and Infection

Endotracheal Intubation in Mice via Direct Laryngoscopy Using an Otoscope

Mice, both wildtype and transgenic, are the principal mammalian model in biomedical research currently. Intubation and mechanical ventilation are necessary for whole animal experiments that require surgery under deep anesthesia or measurements of lung function. Tracheostomy has been the standard for intubating the airway in these mice to allow mechanical ventilation. Orotracheal intubation has been reported but has not been successfully used in many studies because of the substantial technical difficulty or a requirement for highly specialized and expensive equipment. Here we report a technique of direct laryngoscopy using an otoscope fitted with a 2.0 mm speculum and using a 20 G&#160;intravenous catheter as an endotracheal tube. We have used this technique extensively and reliably to intubate and conduct accurate assessments of lung function in mice. This technique has proven safe, with essentially no animal loss in experienced hands. Moreover, this technique can be used for repeated studies of mice in chronic models.

Endotracheal Intubation in Mice via Direct Laryngoscopy Using an Otoscope

We have developed a simple, reliable, and relatively inexpensive method for endotracheal intubation in mice via direct laryngoscopy using an otoscope with a 2.0 mm speculum. This technique is atraumatic and can be used for repeated measurements in chronic experiments. We find it superior to tracheostomy or previously reported nonsurgical techniques.

Mice, both wildtype and transgenic, are the principal mammalian model in biomedical research currently. Intubation and mechanical ventilation are ...

Medicine

In vivo Measurement of the Mouse Pulmonary Endothelial Surface Layer

The endothelial glycocalyx is a layer of proteoglycans and associated glycosaminoglycans lining the vascular lumen. In vivo, the glycocalyx is highly hydrated, forming a substantial endothelial surface layer (ESL) that contributes to the maintenance of endothelial function. As the endothelial glycocalyx is often aberrant in vitro and is lost during standard tissue fixation techniques, study of the ESL requires use of intravital microscopy. To best approximate the complex physiology of the alveolar microvasculature, pulmonary intravital imaging is ideally performed on a freely-moving lung. These preparations, however, typically suffer from extensive motion artifact. We demonstrate how closed-chest intravital microscopy of a freely-moving mouse lung can be used to measure glycocalyx integrity via ESL exclusion of fluorescently-labeled high molecular weight dextrans from the endothelial surface. This non-recovery surgical technique, which requires simultaneous brightfield and fluorescent imaging of the mouse lung, allows for longitudinal observation of the subpleural microvasculature without evidence of inducing confounding lung injury.

In vivo Measurement of the Mouse Pulmonary Endothelial Surface Layer

The endothelial glycocalyx/endothelial surface layer is ideally studied using intravital microscopy. Intravital microscopy is technically challenging in a moving organ such as the lung. We demonstrate how simultaneous brightfield and fluorescent microscopy may be used to estimate endothelial surface layer thickness in a freely-moving in vivo mouse lung.

The endothelial glycocalyx is a layer of proteoglycans and associated glycosaminoglycans lining the vascular lumen. In vivo, the glycocalyx is highly ...

The Utilization of Oropharyngeal Intratracheal PAMP Administration and Bronchoalveolar Lavage to Evaluate the Host Immune Response in Mice

The host immune response to pathogens is a complex biological process. The majority of in vivo studies classically employed to characterize host-pathogen interactions take advantage of intraperitoneal injections of select bacteria or pathogen associated molecular patterns (PAMPs) in mice. While these techniques have yielded tremendous data associated with infectious disease pathobiology, intraperitoneal injection models are not always appropriate for host-pathogen interaction studies in the lung. Utilizing an acute lung inflammation model in mice, it is possible to conduct a high resolution analysis of the host innate immune response utilizing lipopolysaccharide (LPS). Here, we describe the methods to administer LPS using nonsurgical oropharyngeal intratracheal administration, monitor clinical parameters associated with disease pathogenesis, and utilize bronchoalveolar lavage fluid to evaluate the host immune response. The techniques that are described are widely applicable for studying the host innate immune response to a diverse range of PAMPs and pathogens. Likewise, with minor modifications, these techniques can also be applied in studies evaluating allergic airway inflammation and in pharmacological applications.

The host immune response to pathogen infection is a tightly regulated process. Utilizing a lipopolysaccharide lung exposure model in mice, it is possible to conduct high resolution evaluations of the complex mechanisms associated with disease pathogenesis.

The host immune response to pathogens is a complex biological process. The majority of in vivo studies classically employed to characterize host-pathogen ...

A Reversible, Non-invasive Method for Airway Resistance Measurements and Bronchoalveolar Lavage Fluid Sampling in Mice

Airway hyperreactivity (AHR) measurements and bronchoalveolar lavage (BAL) fluid sampling are essential to experimental asthma models, but repeated procedures to obtain such measurements in the same animal are generally not feasible.  Here, we demonstrate protocols for obtaining from mice repeated measurements of AHR and bronchoalveolar lavage fluid samples.  Mice were challenged intranasally seven times over 14 days with a potent allergen or sham treated.  Prior to the initial challenge, and within 24 hours following each intranasal challenge, the same animals were anesthetized, orally intubated and mechanically ventilated.  AHR, assessed by comparing dose response curves of respiratory system resistance (RRS) induced by increasing intravenous doses of acetylcholine (Ach) chloride between sham and allergen-challenged animals, were determined.  Afterwards, and via the same intubation, the left lung was lavaged so that differential enumeration of airway cells could be performed.  These studies reveal that repeated measurements of AHR and BAL fluid collection are possible from the same animals and that maximal airway hyperresponsiveness and airway eosinophilia are achieved within 7-10 days of initiating allergen challenge.  This novel technique significantly reduces the number of mice required for longitudinal experimentation and is applicable to diverse rodent species, disease models and airway physiology instruments.  

Repeated measurements of rodent respiratory physiology and sampling of airway inflammatory cells are desirable, but generally not feasible. Here we describe a repeatable method for orally intubating mice that permits repeated measurements of airway hyperreactivity and sampling of airway inflammatory cells.

Airway hyperreactivity (AHR) measurements and bronchoalveolar lavage (BAL) fluid sampling are essential to experimental asthma models, but repeated ...

Studying Microbial Communities In Vivo: A Model of Host-mediated Interaction Between Candida Albicans and Pseudomonas Aeruginosa in the Airways

Studying host-pathogen interaction enables us to understand the underlying mechanisms of the pathogenicity during microbial infection. The prognosis of the host depends on the involvement of an adapted immune response against the pathogen1. Immune response is complex and results from interaction of the pathogens and several immune or non-immune cellular types2. In vitro studies cannot characterise these interactions and focus on cell-pathogen interactions. Moreover, in the airway3, particularly in patients with suppurative chronic lung disease or in mechanically ventilated patients, polymicrobial communities are present and complicate host-pathogen interaction. Pseudomonas aeruginosa and Candida albicans are both problem pathogens4, frequently isolated from tracheobronchial samples, and associated to severe infections, especially in intensive care unit5. Microbial interactions have been reported between these pathogens in vitro but the clinical impact of these interactions remains unclear6. To study the interactions between C. albicans and P. aeruginosa, a murine model of C. albicans airways colonization, followed by a P. aeruginosa-mediated acute lung infection was performed.

Studying Microbial Communities In Vivo: A Model of Host-mediated Interaction Between Candida Albicans and Pseudomonas Aeruginosa in the Airways

While in vitro study of host-pathogen interactions allow the characterization of specific immune responses, in vivo models are required to observe the effects of complex responses. Using Candida albicans exposure followed by Pseudomonas aeruginosa-mediated lung infection, we established a murine model of microbial interactions involved in ventilator-associated pneumonia pathogenicity.

Studying host-pathogen interaction enables us to understand the underlying mechanisms of the pathogenicity during microbial infection. The prognosis of ...

Evaluation of Respiratory System Mechanics in Mice using the Forced Oscillation Technique

The forced oscillation technique (FOT) is a powerful, integrative and translational tool permitting the experimental assessment of lung function in mice in a comprehensive, detailed, precise and reproducible manner. It provides measurements of respiratory system mechanics through the analysis of pressure and volume signals acquired in reaction to predefined, small amplitude, oscillatory airflow waveforms, which are typically applied at the subject's airway opening. The present protocol details the steps required to adequately execute forced oscillation measurements in mice using a computer-controlled piston ventilator (flexiVent; SCIREQ Inc, Montreal, Qc, Canada). The description is divided into four parts: preparatory steps, mechanical ventilation, lung function measurements, and data analysis. It also includes details of how to assess airway responsiveness to inhaled methacholine in anesthetized mice, a common application of this technique which also extends to other outcomes and various lung pathologies. Measurements obtained in na&iuml;ve mice as well as from an oxidative-stress driven model of airway damage are presented to illustrate how this tool can contribute to a better characterization and understanding of studied physiological changes or disease models as well as to applications in new research areas.

The present protocol provides a detailed step-by-step description of the procedures required to execute measurements of respiratory system mechanics as well as the assessment of airway responsiveness to inhaled methacholine in mice using the forced oscillation technique (flexiVent; SCIREQ Inc, Montreal, Qc, Canada).

The  forced oscillation technique (FOT) is a powerful, integrative and translational  tool permitting the experimental assessment of lung function in mice ...

Assessment of Respiratory Function in Conscious Mice by Double-chamber Plethysmography

Air volume changes created by a conscious subject breathing spontaneously within a body box are at the basis of plethysmography, a technique used to non-invasively assess some features of the respiratory function in humans as well as in laboratory animals. The present article focuses on the application of the double-chamber plethysmography (DCP) in small animals. It provides background information on the methodology as well as a detailed step-by-step procedure to successfully assess respiratory function in conscious, spontaneously breathing animals in a non-invasive manner. The DCP can be used to monitor the respiratory function of multiple animals in parallel, as well as to identify changes induced by aerosolized substances over a chosen time period and in a repeated manner. Experiments on control and allergic mice are used herein to demonstrate the utility of the technique, explain the associated outcome parameters, as well as to discuss the related advantages and shortcomings. Overall, the DCP provides valid and theoretically sound readouts that can be trusted to evaluate the respiratory function of conscious small animals both at baseline and after challenges with aerosolized substances.

The objective of the present article is to provide a detailed description of the recommended procedures to evaluate respiratory function in conscious mice by double-chamber plethysmography.

Air volume changes created by a conscious subject breathing spontaneously within a body box are at the basis of plethysmography, a technique used to ...

Ex vivo Method for High Resolution Imaging of Cilia Motility in Rodent Airway Epithelia

An ex vivo technique for imaging mouse airway epithelia for quantitative analysis of motile cilia function important for insight into mucociliary clearance function has been established. Freshly harvested mouse trachea is cut longitudinally through the trachealis muscle and mounted in a shallow walled chamber on a glass-bottomed dish. The trachea sample is positioned along its long axis to take advantage of the trachealis muscle to curl longitudinally. This allows imaging of ciliary motion in the profile view along the entire tracheal length. Videos at 200 frames/sec are obtained using differential interference contrast microscopy and a high speed digital camera to allow quantitative analysis of cilia beat frequency and ciliary waveform. With the addition of fluorescent beads during imaging, cilia generated fluid flow also can be determined. The protocol time spans approximately 30 min, with 5 min for chamber preparation, 5-10 min for sample mounting, and 10-15 min for videomicroscopy.

Ex vivo Method for High Resolution Imaging of Cilia Motility in Rodent Airway Epithelia

An easy and reliable technique for visualizing and quantifying airway cilia motility and cilia generated flow using mouse trachea is described. This technique can be modified to determine how a wide range of factors influence cilia motility, including pharmacological agents, genetic factors, environmental exposures, and/or mechanical factors such as mucus load.

An ex vivo technique for imaging mouse airway epithelia for quantitative  analysis of motile cilia function important for insight into mucociliary  ...

Nasal Potential Difference to Quantify Trans-epithelial Ion Transport in Mice

The nasal potential difference test has been used for almost three decades to assist in the diagnosis of cystic fibrosis (CF). It has proven to be helpful in cases of attenuated, oligo- or mono-symptomatic forms of CF usually diagnosed later in life, and of CF-related disorders such as congenital bilateral absence of vas deferens, idiopathic chronic pancreatitis, allergic bronchopulmonary aspergillosis, and bronchiectasis. In both clinical and preclinical settings, the test has been used as a biomarker to quantify responses to targeted therapeutic strategies for CF. Adapting the test to a mouse is challenging and can entail an associated mortality. This paper describes the adequate depth of anesthesia required to maintain a nasal catheter in situ for continuous perfusion. It lists measures to avoid broncho-aspiration of solutions perfused in the nose. It also describes the animal care at the end of the test, including administration of a combination of antidotes of the anesthetic drugs, leading to rapidly reversing the anesthesia with full recovery of the animals. Representative data obtained from a CF and a wild-type mouse show that the test discriminates between CF and non-CF. Altogether, the protocol described here allows reliable measurements of the functional status of trans-epithelial chloride and sodium transporters in spontaneously breathing mice, as well as multiple tests in the same animal while reducing test-related mortality.

Here, we present a protocol to measure nasal potential difference in mice. The test quantifies the function of transmembrane ion transporters such as the cystic fibrosis transmembrane conductance regulator and the epithelial sodium channel. It is valuable to evaluate the efficacy of novel therapies for cystic fibrosis.

The nasal potential difference test has been used for almost three decades to assist in the diagnosis of cystic fibrosis (CF). It has proven to be helpful ...

In vivo Evaluation of Mucociliary Clearance in Mice

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Murine Model of Allergen Induced Asthma

Endotracheal Intubation in Mice via Direct Laryngoscopy Using an Otoscope

In vivo Measurement of the Mouse Pulmonary Endothelial Surface Layer

The Utilization of Oropharyngeal Intratracheal PAMP Administration and Bronchoalveolar Lavage to Evaluate the Host Immune Response in Mice

A Reversible, Non-invasive Method for Airway Resistance Measurements and Bronchoalveolar Lavage Fluid Sampling in Mice

Studying Microbial Communities In Vivo: A Model of Host-mediated Interaction Between Candida Albicans and Pseudomonas Aeruginosa in the Airways

Evaluation of Respiratory System Mechanics in Mice using the Forced Oscillation Technique

Assessment of Respiratory Function in Conscious Mice by Double-chamber Plethysmography

Ex vivo Method for High Resolution Imaging of Cilia Motility in Rodent Airway Epithelia

Nasal Potential Difference to Quantify Trans-epithelial Ion Transport in Mice