Name: Ex vivo Method for High Resolution Imaging of Cilia Motility in Rodent Airway Epithelia (Video) | JoVE Journal
Uploaded: 2013-08-08T13:00:00.000+00:00
Duration: 8 min 25 s
Description: Learn an ex vivo method for high-resolution imaging of cilia motility in rodent airway epithelia using freshly isolated mouse trachea. This protocol enables quantitative assessment of ciliary beat frequency and mucociliary clearance.

ai_agent_IP

ai_agent

maintenance

instantTrialDurationMinutes

isBiopharma

enableNewMyJoveSectionDesign

educationFeatures

seoMetaApi

ai_copilot_institution_blacklist

ai_chat_institution_blacklist

freeAccessWindow

ssoRoleCapture

scheduledMaintenanceBanner

joveLabsFeatures

disable_autoplay_for_bots

extended_trial_duration

5 minutes

playerTrialDesktop

The project was supported by NIH grant U01HL098180 from the National Heart, Lung, and Blood Institute. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Heart, Lung, and Blood Institute or the National Institutes of Health.

1. Reagents Setup
1.1 Dissection and Imaging Medium
Leibovitz's L-15 medium (L15) is supplemented with FBS (10%) and Penicillin-Streptomycin (100 units/ml of penicillin G sodium and 100 &mu;g/ml of streptomycin sulfate) is used during both harvest and imaging of trachea samples.
2. Shallow Walled Culture Chamber Assembly
The chamber used to hold trachea tissue is shown in Figure 1. The floor of the chamber is the glass bottomed 35 mm culture dish. The top and side of the chamber is generated by placing a small piece of 0.3 mm thick silicone sheeting over the glass dish. The edge of the sheet is cut to fit the round bottom dish and the center is further trimmed to form a shallow central chamber (see steps 2.1-2.3). On top of this assembly, a 18 mm round glass cover slip is placed to generate an enclosed chamber suitable for imaging.
<ol>
	<li>Completely cover a 18 mm round glass cover slip with a square of 0.3 mm thick silicone sheeting.</li>
	<li>With standard sharp dissection scissors trim off the excess silicone sheeting from around the edge of the cover slip (resulting in a circular layer of silicone sheeting).</li>
	<li>With a sharp scalpel cut out and remove a central square of silicone sheeting (approximately 10 mm square) from the middle of the covered cover slip (thus forming the top and sides of the imaging chamber).</li>
</ol>
3. Trachea Harvest and Preparation
<ol>
	<li>Euthanize mice in accordance with local institutional animal care and use committee and/or governmental guidelines and remove trachea into a standard plastic 35 mm culture dish with enough L15 media to submerge tissue (authors have used mice just before birth at gestation day 16.5, through to newborn, neonatal, and adult animals 2).</li>
	<li>Remove non-trachea associated tissue attached to the outside of the trachea using blunt dissection.</li>
	<li>Remove the larynx with a transverse cut just below the cricoid cartilage using micro dissection scissors (Figure 2A, line 1).</li>
	<li>Remove the left and right primary bronchial branches with a transverse cut just above the carina using micro dissection scissors (Figure 2A, line 2).</li>
	<li>Gently hold the trachea with fine forceps and flush the lumen 2-3 times with ~1 ml of the bathing L15 medium with a P1000 Pipetman and P1000 tip to clear any mucus and blood.</li>
	<li>Cut out a 3-4 ring segment of trachea using a transverse cut with micro dissection scissors (Figure 2B, line 1).</li>
	<li>Cut longitudinally through the middle of the trachealis muscle of this short trachea segment using micro dissection scissors (Figure 2B, line 2).</li>
	<li>Cut longitudinally through the middle of the tracheal cartilage rings using micro dissection scissors (Figure 2C, line), leaving two sections of trachea tissue suitable for imaging (single section shown in Figure 2D).</li>
</ol>
4. Cilia Imaging
<ol>
	<li>Transfer trachea tissue segments, lumen side down, onto the middle of a 35 mm glass bottom culture dish with 100-200 &mu;l of L-15 medium.</li>
	<li>To quantify cilia generated flow add a small amount of fluorescent 0.20 &mu;m microspheres to the L-15 medium bathing the trachea tissue sample (~1 x 1011 particles/ml or 20 &mu;l/ml of the Fluoresbrite 2.5% aqueous microsphere suspension).</li>
	<li>Place the imaging chamber prepared previously (See Procedure 1-3 above) on top of the trachea sample silicone sheet side down, with the trachea sample located in the middle of the shallow walled chamber formed by the window cut into the silicone sheeting (Figure 1).</li>
	<li>Gently push down on the coverslip to secure in place and remove any excess L-15 media from the edges using a P1000 tip and P1000 Pipetman.</li>
	<li>Mount the 35 mm glass bottom culture dish and trachea sample on a inverted microscope with 100X objective and DIC optics.</li>
	<li>Using a high-speed camera and movie acquisition software collect movies of cilia motility at 200+ frames per second along the ciliated tracheal epithelia of the curled up edge of the trachealis muscle (Figure 2D Outlined box, E: example still image from recorded movie)(Supplementary Movie 1).</li>
	<li>Using FITC epifluorescent imaging and a low light CCD camera (See equipment list above), collect movies of fluorescent microsphere movement across the surface of the ciliated tracheal epithelia to allow later quantification of cilia generated flow (Supplementary Movie 2).</li>
</ol>
5. Quantitation of Cilia Motility
<ol>
	<li>Load DIC imaged movies into the ImageJ software package and following the technique described by Iain Drummond 13 use the line tool to draw a raster line crossing the beating cilia. A "reslice" of this line generates a kymograph type image where cilia movement across the line generates a wave pattern. The number of pixels between each wave peak is them measured (one pixel = one movie frame) from which the number of beats per minute (i.e. Hz) can then be calculated.</li>
	<li>To measure cilia generated flow load fluorescent bead movies into the ImageJ software package, and use the MTrackJ plugin 14 to manually track fluorescent beads across the surface of the trachea epithelia and let the software generate bead velocity and directionality for each bead tracked.</li>
</ol>
Care should be taken when using fluorescent beads to quantify flow as the distance from the beating cilia can strongly influence measured flow magnitude. The bead movement in Supplemental Movie 2 is a good example of this. In this example bead movement is fast at the surface of the beating cilia and significantly drops off for beads further away in the bathing media. To control for this, bead tracings should only be collected at a fixed distance above the ciliated epithelia in all samples to allow correct comparisons to be made. Finally, to obtain representative data on fluid flow, beads should be tracked for as long as possible; we prefer to use bead tracks that cover 10+ ciliated cells for calculating fluid flow.

<ol><li>Stannard, W., O'Callaghan, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Ciliary+function+and+the+role+of+cilia+in+clearance%5BTitle%5D)+AND+%22J.+Aerosol.+Med%22%5BJournal%5D)">Ciliary function and the role of cilia in clearance.</a> J. Aerosol. Med. 19, 110-115 (2006).</li>
<li>Francis, R. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+initiation+and+maturation+of+cilia+generated+flow+in+the+newborn+and+postnatal+mouse+airway%5BTitle%5D)+AND+%22American+Journal+of+Physiology%3A+Lung+Cellular+and+Molecular+Physiology%22%5BJournal%5D)">The initiation and maturation of cilia generated flow in the newborn and postnatal mouse airway.</a> American Journal of Physiology: Lung Cellular and Molecular Physiology. 296, 1067-1075 (2009).</li>
<li>Aune, C. N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Mouse+model+of+heterotaxy+with+single+ventricle+spectrum+of+cardiac+anomalies%5BTitle%5D)+AND+%22Pediatric+Research%22%5BJournal%5D)">Mouse model of heterotaxy with single ventricle spectrum of cardiac anomalies.</a> Pediatric Research. 63, 9-14 (2008).</li>
<li>Tan, S. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Heterotaxy+and+complex+structural+heart+defects+in+a+mutant+mouse+model+of+primary+ciliary+dyskinesia%5BTitle%5D)+AND+%22J.+Clin.+Invest%22%5BJournal%5D)">Heterotaxy and complex structural heart defects in a mutant mouse model of primary ciliary dyskinesia.</a> J. Clin. Invest. 117, 3742-3752 (2007).</li>
<li>Zhang, Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Massively+parallel+sequencing+identifies+the+gene+Megf8+with+ENU-induced+mutation+causing+heterotaxy%5BTitle%5D)+AND+%22Proc.+Natl.+Acad.+Sci.+U.S.A%22%5BJournal%5D)">Massively parallel sequencing identifies the gene Megf8 with ENU-induced mutation causing heterotaxy.</a> Proc. Natl. Acad. Sci. U.S.A. 106, 3219-3224 (2009).</li>
<li>Caruso, G., Gelardi, M., Passali, G. C., de Santi, M. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Nasal+scraping+in+diagnosing+ciliary+dyskinesia%5BTitle%5D)+AND+%22Am.+J.+Rhinol%22%5BJournal%5D)">Nasal scraping in diagnosing ciliary dyskinesia.</a> Am. J. Rhinol. 21, 702-705 (2007).</li>
<li>Leigh, M. W., Zariwala, M. A., Knowles, M. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Primary+ciliary+dyskinesia%3A+improving+the+diagnostic+approach%5BTitle%5D)+AND+%22Curr.+Opin.+Pediatr%22%5BJournal%5D)">Primary ciliary dyskinesia: improving the diagnostic approach.</a> Curr. Opin. Pediatr. 21, 320-325 (2009).</li>
<li>Delmotte, P., Sanderson, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Ciliary+beat+frequency+is+maintained+at+a+maximal+rate+in+the+small+airways+of+mouse+lung+slices%5BTitle%5D)+AND+%22Am.+J.+Respir.+Cell+Mol.+Biol%22%5BJournal%5D)">Ciliary beat frequency is maintained at a maximal rate in the small airways of mouse lung slices.</a> Am. J. Respir. Cell Mol. Biol. 35, 110-117 (2006).</li>
<li>Choe, M. M., Tomei, A. A., Swartz, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Physiological+3D+tissue+model+of+the+airway+wall+and+mucosa%5BTitle%5D)+AND+%22Nat.+Protoc%22%5BJournal%5D)">Physiological 3D tissue model of the airway wall and mucosa.</a> Nat. Protoc. 1, 357-362 (2006).</li>
<li>Fulcher, M. L., Gabriel, S., Burns, K. A., Yankaskas, J. R., Randell, S. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Well-differentiated+human+airway+epithelial+cell+cultures%5BTitle%5D)+AND+%22Methods+Mol.+Med%22%5BJournal%5D)">Well-differentiated human airway epithelial cell cultures.</a> Methods Mol. Med. 107, 183-206 (2005).</li>
<li>You, Y., Richer, E. J., Huang, T., Brody, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Growth+and+differentiation+of+mouse+tracheal+epithelial+cells%3A+selection+of+a+proliferative+population%5BTitle%5D)+AND+%22Am.+J.+Physiol.+Lung+Cell+Mol.+Physiol%22%5BJournal%5D)">Growth and differentiation of mouse tracheal epithelial cells: selection of a proliferative population.</a> Am. J. Physiol. Lung Cell Mol. Physiol. 283, 1315-1321 (2002).</li>
<li>Thomas, B., Rutman, A., O'Callaghan, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Disrupted+ciliated+epithelium+shows+slower+ciliary+beat+frequency+and+increased+dyskinesia%5BTitle%5D)+AND+%22Eur.+Respir.+J%22%5BJournal%5D)">Disrupted ciliated epithelium shows slower ciliary beat frequency and increased dyskinesia.</a> Eur. Respir. J. 34, 401-404 (2009).</li>
<li>Drummond, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Studying+cilia+in+zebrafish%5BTitle%5D)+AND+%22Methods+Cell+Biol%22%5BJournal%5D)">Studying cilia in zebrafish.</a> Methods Cell Biol. 93 (08), 197-217 (2009).</li>
<li>Meijering, E., Dzyubachyk, O., Smal, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Methods+for+cell+and+particle+tracking%5BTitle%5D)+AND+%22Methods+Enzymol%22%5BJournal%5D)">Methods for cell and particle tracking.</a> Methods Enzymol. 504, 183-200 (2012).</li>
<li>Sisson, J. H., Stoner, J. A., Ammons, B. A., Wyatt, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((All-digital+image+capture+and+whole-field+analysis+of+ciliary+beat+frequency%5BTitle%5D)+AND+%22J.+Microsc%22%5BJournal%5D)">All-digital image capture and whole-field analysis of ciliary beat frequency.</a> J. Microsc. 211, 103-111 (2003).</li>
<li>Schwabe, G. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Primary+ciliary+dyskinesia+associated+with+normal+axoneme+ultrastructure+is+caused+by+DNAH11+mutations%5BTitle%5D)+AND+%22Hum.+Mutat%22%5BJournal%5D)">Primary ciliary dyskinesia associated with normal axoneme ultrastructure is caused by DNAH11 mutations.</a> Hum. Mutat. 29, 289-298 (2008).</li>
<li>Shah, A. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Loss+of+Bardet-Biedl+syndrome+proteins+alters+the+morphology+and+function+of+motile+cilia+in+airway+epithelia%5BTitle%5D)+AND+%22Proc.+Natl.+Acad.+Sci.+U.S.A%22%5BJournal%5D)">Loss of Bardet-Biedl syndrome proteins alters the morphology and function of motile cilia in airway epithelia.</a> Proc. Natl. Acad. Sci. U.S.A. 105, 3380-3385 (2008).</li>
<li>Clary-Meinesz, C. F., Cosson, J., Huitorel, P., Blaive, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Temperature+effect+on+the+ciliary+beat+frequency+of+human+nasal+and+tracheal+ciliated+cells%5BTitle%5D)+AND+%22Biology+of+the+Cell+%2F+under+the+auspices+of+the+European+Cell+Biology+Organization%22%5BJournal%5D)">Temperature effect on the ciliary beat frequency of human nasal and tracheal ciliated cells.</a> Biology of the Cell / under the auspices of the European Cell Biology Organization. 76, 335-338 (1992).</li>
</ol>

Authors have nothing to disclose.

Measurement of cilia beat frequency (CBF) is relatively easy using high power microscope objectives and fast image acquisition hardware 13,15, and explains why CBF measurements form the basis of most studies investigating mucociliary clearance during health and disease. However, while CBF measurement is essential for understanding mucociliary clearance, measurement of CBF alone ignores the underlying importance of both ciliary generated flow and cilia beat waveform, both of which are more difficult to measure,...

Analysis of motile cilia function in the airway epithelia is experimentally important for elucidating the genetic and environmental factors that can affect mucociliary clearance and pulmonary health 1. The simple protocol developed for imaging the mouse airway epithelia provides an efficient method to interrogate airway cilia motility in mutant and knockout mouse models and require only basic skills in mouse tracheal tissue dissection and ex vivo imaging of airway cilia motility with high resolution videomicroscopy. This protocol was established and refined during a large-scale mouse mutagenesis screen to allow rapid evaluation of motile cilia function (cilia beat frequency, cilia beat shape, cilia generated flow) in mutants with congenital heart disease associated with heterotaxy 2-5.
Current techniques used to study airway cilia motility can be grouped into either the acute ex vivo type or longer term in vitro experimental approaches. Acute experiments include ex vivo visualization of human nasal/airway brush biopsies 6,7 and analysis of simple transverse airway sections 8. The in vitro approaches utilize various cell culture techniques to generate sheets of differentiated ciliated epithelia such as in air liquid interface cultures or airway suspension cultures9-11. However, these airway epithelia reciliation techniques require very significant investment in time and training before any useable ciliated epithelial cells are produced for experimentation (4-6 weeks 9,10). While acute ex vivo analysis of airway epithelial brush biopsies are commonly used for human clinical studies, this method is not usable in mouse studies due to exacerbated mechanical tissue injury 12.
 
The technique outlined in this protocol for analysis of mouse tracheal airway epithelia is not only simple to execute, but it requires no special dissection skills nor any specialized equipment besides those standard for imaging by videomicroscopy. There are many advantages to this simple protocol. First, as the mouse trachea tissue harvest is quick and easy to perform, it allows for rapid assessment of airway cilia function in a large number of mice. This can include acute analysis of the short term effects of different in vitro treatments. Second, being an ex vivo technique, the ciliated airway epithelium remains attached to it underlying supporting tissues and thus retain associated cell signaling pathways. Therefore in comparison to in vitro reciliated airway epithelia, this preparation is a better representation of the natural in vivo tissue environment. Third, this protocol allows the acquisition of a number of different quantitative parameters that can provide the objective assessment of motile cilia function. Finally, in contrast to other current methods for airway cilia visualization, this protocol allows for visualization of the cilia at right angles to the cilia beat direction, allowing profile view of the cilia that is optimal for high resolution imaging of cilia beat and metachronal wave generation.
This protocol can be modified in a number of ways to address a wide range of experimental needs such as the role of pharmacological agents, genetic factors, environmental exposures, and/or mechanical factors such as mucus load on airway cilia function and generation/maintenance of airway cilia beat and metachronal wave propagation.

<table><tbody><tr><td>Leibovitz&amp;rsquo;s L-15 Medium </td><td>Invitrogen</td><td>21083-027</td><td>No phenol red </td></tr><tr><td>Fetal bovine serum </td><td>Hyclone</td><td>SH30088.03</td><td></td></tr><tr><td>Penicillin-Streptomycin </td><td>Invitrogen</td><td>15140-122</td><td></td></tr><tr><td>2x fine forceps </td><td>Roboz</td><td>RS-4976</td><td></td></tr><tr><td>Dissection scissors </td><td>Roboz</td><td>RS-5676</td><td></td></tr><tr><td>Micro dissection scissors </td><td>Roboz</td><td>RS-5620</td><td></td></tr><tr><td>Scalpel </td><td>Roboz</td><td>RS-9801-15</td><td></td></tr><tr><td>P1000 pipetman </td><td>Gilson, Inc</td><td>F123602</td><td></td></tr><tr><td>P1000 tips </td><td>Molecular BioProducts</td><td>2079E</td><td></td></tr><tr><td>18 mm round glass cover slips </td><td>Fisher Scientific</td><td>430588</td><td></td></tr><tr><td>Plastic 35 mm culture dishes </td><td>Corning</td><td>430588</td><td></td></tr><tr><td>Glass bottom 35 mm culture dishes </td><td>Warner Instruments</td><td>W3 64-0758</td><td></td></tr><tr><td>Silicone sheet 0.012&quot; (0.3 mm) thick </td><td>AAA Acme Rubber Co</td><td>CASS-.012X36-63908</td><td></td></tr><tr><td>0.20 &amp;mu;m diameter Fluoresbrite YG Carboxylate Microspheres </td><td>Polysciences</td><td>09834-10</td><td></td></tr><tr><td>Inverted microscope, with 100x oil objective and DIC filters </td><td>Lecia</td><td>DMIRE2</td><td>Brand is not critical. </td></tr><tr><td>100-watt mercury lamp, epifluorescent FITC excitation/emission filters</td><td>Lecia</td><td></td><td>Brand is not critical.</td></tr><tr><td>Microscope stage Incubator</td><td>Lecia</td><td>11521749</td><td>Not required if imaging cilia at room temperature</td></tr><tr><td>High-speed camera bright field </td><td>Vision Research</td><td>Phantom v4.2</td><td>Brand is not critical. Must be faster than 125 fps</td></tr><tr><td>High-speed fluorescent camera </td><td>Hamamatsu</td><td>C9100-12</td><td>Brand is not critical. Must be faster than 10 fps</td></tr><tr><td>Movie analysis software</td><td>National Institutes of Health</td><td>ImageJ with MtrackJ plugin</td><td></td></tr></tbody></table>

ex vivo method for high resolution imaging of cilia motility in rodent airway epithelia

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.

Control airway cilia should be clearly visible and seen to beat in a coordinated manner (Supplemental Movie 1; movie playback is slowed to 15% real time), with noticeable flow in the direction of cilia beat (Supplemental Movie 2; movie playback is 100% real time). Quantitation of cilia movies should yield results similar to that seen in Figure 3. Collection of high-speed DIC movies makes possible the quantification of cilia beat frequency (Figure 3C) and...

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  ...

The overall goal of this procedure is to visualize and quantify respiratory CLIA motility and CLIA generated flow using freshly isolated mouse trachea. This is accomplished by first preparing a shallow imaging chamber. The next steps of the procedure are to harvest and mount the tracheal tissue for imaging.It is then shown how to visualize and record C motility and CLIA generated flow using an inverted microscope with high speed videography through a 100 XDIC objective. The final step is to analyze the video by quantifying CLIA beat frequency and CLIA generated flow using Image J software. Ultimately, the results can show changes in respiratory CLIA motility and CLIA generated flow through high speed video microscopy.The main advantage of this technique over existing methods is that it allows reproducible imaging of the rodent tra epithelium for consistent quantitative assessments of ciliary motion. This can be accomplished within 20 to 30 minutes with minimal training. This method can help address key questions relating to the cell and molecular mechanisms regulating muci clearance in the eWAY, especially when it is used in combination with different genetically engineered and mut

ex-vivo-method-for-high-resolution-imaging-cilia-motility-rodent

Research

JoVE Journal

Biology

Here we develop the tools necessary for ex vivo live imaging to trace single cell divisions in the mouse E8.5...

Ex vivo Live Imaging of Single Cell Divisions in Mouse Neuroepithelium

Cilia of olfactory sensory neurons contain proteins of the signal transduction cascade, but a detailed spatial analysis of their distribution is difficult in cryosections. We describe here an optimized approach for whole mount labeling and en face visualization of ciliary proteins.

Whole Mount Labeling of Cilia in the Main Olfactory System of Mice

Using high-resolution differential interference contrast (DIC) microscopy, an ex vivo observation of the beating of motile ependymal cilia located within the mouse brain ventricles is demonstrated by live-imaging. The technique allows a recording of the unique ciliary beating frequency and beating angle as well as their intracellular calcium oscillation pacing properties.

Live Imaging of the Ependymal Cilia in the Lateral Ventricles of the Mouse Brain

In this study, a detailed light microscopic technique was optimized for real-time observation and analysis of the motion of CPEC cilia ex vivo together with an electron microscopic method for ultrastructural analysis.

Observation of the Ciliary Movement of Choroid Plexus Epithelial Cells Ex Vivo

We describe a method to visualize GFP-labeled γδ IELs using intravital imaging of murine small intestine by inverted spinning disk confocal microscopy. This technique enables the tracking of live cells within the mucosa for up to 4 h and can be used to investigate a variety of intestinal immune-epithelial interactions.

Intravital Imaging of Intraepithelial Lymphocytes in Murine Small Intestine

We describe a simple protocol to develop mucociliary epithelial organoids from deep ectoderm cells isolated from Xenopus laevis embryos. The multipotent progenitors regenerate epithelial goblet cell precursors and allow live tracking of the initiation and progression of the cell transitions on the surface of organoids.

Mucociliary Epithelial Organoids from Xenopus Embryonic Cells: Generation, Culture and High-Resolution Live Imaging

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.

In vivo Evaluation of Mucociliary Clearance in Mice

To guarantee a successful and high-quality ciliary functional analysis for PCD diagnosis, a precise and careful method for respiratory epithelium sampling and processing is essential. To continue providing PCD diagnostic service during the COVID-19 pandemic, the ciliary videomicroscopy protocol has been updated to include appropriate infection control measures.

Nasal Brushing Sampling and Processing Using Digital High Speed Ciliary Videomicroscopy &ndash; Adaptation for the COVID-19 Pandemic

This protocol describes nasal epithelial cell collection, expansion, and differentiation to organotypic airway epithelial cell models and quantification of cilia beat frequency via live-cell imaging and custom-built scripts.

Collection, Expansion, and Differentiation of Primary Human Nasal Epithelial Cell Models for Quantification of Cilia Beat Frequency

Air-liquid interface culture is commonly used to develop pseudostratified airway epithelium by differentiating primary normal human bronchial epithelial cells that mimic the apical side of the lung airway. Here, we describe an easy protocol for determining its quality by monitoring its biophysical properties, such as ciliary function and membrane integrity.

Determining Ciliary Function and Membrane Impermeability of the Pseudostratified Lung Airway Epithelium

This method describes the exploitation of the endogenous signal in an ARL13B-CENTRIN-2 transgenic mouse model to image primary cilia in situ, in three dimensions across large areas of mineralized tissue, from tissue collection to analysis of cilia organization. This protocol may also be applied to other types of tissues.

Volumetric Imaging and Analysis of Primary Cilia in Musculoskeletal Tissue using the ARL13B-CENTRIN-2 Mouse Model

Learn an ex vivo method for high-resolution imaging of cilia motility in rodent airway epithelia using freshly isolated mouse trachea. This protocol enables quantitative assessment of ciliary beat frequency and mucociliary clearance.

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

Explore More Videos