We thank Adam Chicoine at the Brigham and Women&#8217;s Human Immunology Center Flow Core for his help with flow cytometric sorting. This work was supported by National Institutes of Health Grants R01 HL120952 (N.A.B.), R01 AI134989 (N.A.B), U19 AI095219 (N.A.B., L.G.B), and K08 AI132723 (L.G.B), and by the American Academy of Allergy, Asthma, and Immunology (AAAAI)/ American Lung Allergic Respiratory Disease Award (N.A.B.), by the AAAAI Foundation Faculty Development Award (L.G.B.), by the Steven and Judy Kaye Young Innovators Award (N.A.B.), by the Joycelyn C. Austen Fund for Career Development of Women Physician Scientists (L.G.B.), and by a generous donation by the Vinik family (L.G.B.).

Before conducting the following experiments, ensure that all animal care use and protocols are approved by the Institutional Animal Care and Use Committee (IACUC) and performed in accord with the National Research Council&#39;s &#34;Guide for the Care and Use of Laboratory Animals&#34;&#160;(8th&#160;Edition, 2011) and the ARRIVE guidelines.&#160;All procedures described below have been reviewed and approved by the Institutional Animal Care and Use Committee at the Brigham and Women&#39;s Hospital.
1. Preparation of Reagents
<ol>
	<li>Prepare Dispase digestion solution, which is a PBS solution containing 16 U/mL dispase and 20 &#181;g/mL DNase I. Ensure that the dispase powder is fully dissolved before warming up the solution in a water bath at 37 &#176;C.</li>
	<li>Add 5% heat-inactivated fetal bovine serum (FBS) to Dulbecco Modified Eagle Medium (DMEM) to make a stopping solution.</li>
	<li>Prepare Tyrode I buffer: add 26 U/mL papain (20 &#181;L/mL of 48 U/mg papain solution) and 10 &#181;L/mL L-cysteine to HEPES-Tyrode&#39;s buffer without calcium.</li>
	<li>Prepare Tyrode II buffer: add 2 &#956;L/mL leupeptin (5 mg/mL) to HEPES-Tyrode&#39;s buffer with calcium.</li>
	<li>Prepare FACS buffer: use Hanks&#39;&#160;Balanced Salt Solution (HBSS) without calcium, magnesium &#38; phenol red, supplemented with 2 mM ethylenediaminetetraacetic acid (EDTA) and add 2% FBS.</li>
</ol>
2. Dissection of Mouse Trachea
NOTE: Mice used in this protocol are ChAT(BAC)-eGFP (B6.Cg-Tg(RP23- 268L19-EGFP)2Mik/J), 3-6 months of age of both sexes. Minimize the exposure of the tissue to direct light to reduce photobleaching of eGFP.
<ol>
	<li>Euthanize the mouse with 100 mg/kg pentobarbital euthanasia solution injected intraperitoneally or using standard protocols approved by the IACUC.</li>
	<li>Fix the mouse on a surgical board in the supine position with 21 G needles with extended upper and lower extremities. Spray the fur with 70% EtOH to sanitize the area.</li>
	<li>Using straight forceps, lift the skin and fur of the abdomen and make an incision in the center with dissecting scissors (straight scissors, 3 cm). Using the scissors, separate the skin from the subcutaneous tissue from the abdomen to the mandibula. While holding the subcutaneous tissue up with the forceps, make a small incision with the scissors in the center of the abdominal wall.</li>
	<li>Open the peritoneum with a V-shaped incision. Using the forceps, gently move the small intestine to the side, locate the abdominal aorta and vena cava and make an incision with the dissecting scissors to allow for rapid exsanguination.</li>
	<li>Locate the diaphragm. Using an 18 G needle, make an opening in the diaphragm just below the sternum to deflate the lungs. Carefully separate the diaphragm from the rib cage using sharp-pointed straight dissecting scissors to cut along the base of the ribs.</li>
	<li>Using the forceps, lift the exposed end of the sternum and cut the sternum longitudinally from the base of the rib cage to the neck. Make a central cervical incision with short straight scissors (2 cm) and separate the two lobes of the submandibular gland.</li>
	<li>Carefully remove the surrounding connective tissue and the thymus overlying the carina with a pair of fine point high precision forceps.</li>
	<li>Dissect the trachea free first by separating the proximal end at the level of the epiglottis and then by dissecting the distal end at the level of the bifurcation of the trachea.</li>
	<li>Locate the epiglottis and cut the trachea longitudinally from the epiglottis to the carina.</li>
</ol>
3. Tracheal Epithelial Digestion
<ol>
	<li>Place the trachea into a 1.5 mL tube containing 750 &#181;L of pre-warmed (to 37 &#176;C) dispase digestion solution. Incubate on a shaker at 200 rpm for 40 min at room temperature. Cover the tube with aluminum foil to reduce the exposure to direct light.</li>
	<li>Add 750 &#181;L of cold DMEM with 5% FBS to stop the reaction. Place on ice.</li>
	<li>Transfer the trachea to a Petri dish (100 mm x 15 mm) and place under a dissecting microscope. Orient the trachea with the epithelial side facing up. The longitudinally dissected trachea has a semi-cylindrical shape maintained by the cartilaginous rings. The epithelium is on the concave surface.</li>
	<li>Tether the epiglottis area of the trachea with straight forceps to the Petri dish and using a size 22 disposable scalpel, scrape the epithelium off the trachea. The epithelial layer separates as a translucent sheet.</li>
	<li>Mince the epithelium with the scalpel. Transfer the epithelial layer to a 2 mL tube.</li>
	<li>Rinse the Petri dish with 750 &#181;L of Tyrode I buffer and transfer to the 2 mL tube containing the epithelial layer.</li>
	<li>Incubate the tracheal epithelial layer in Tyrode I buffer for 30 min at 37 &#176;C on a shaker at 200 rpm. Cover the tube with aluminum foil to reduce the exposure to direct light.</li>
	<li>Add 750 &#181;L of cold Tyrode II buffer. Vortex the digested tissue vigorously for 20-30 s. Triturate the homogenate with a syringe attached to an 18 G needle 10 times.&#160; Switch to a 21 G gauge needle and triturate 10-20 more times.</li>
	<li>Filter the cells through a 100 &#181;m strainer into a 50 mL conical tube. Add 30:1 vol/vol of cold FACS buffer.</li>
	<li>Spin at 350 x g for 10 min at 4 &#176;C and discard the supernatant. Resuspend the pellet in cold FACS buffer and transfer the suspension to a 12 mm x 75 mm (5 mL) polystyrene tube. Spin again at 350 x g for 10 min at 4 &#176;C and discard the supernatant.</li>
	<li>Re-suspend the pellet in 100 &#181;L of FACS buffer.</li>
	<li>Add 1 &#181;L of anti-mouse CD16/32 blocking antibody to block non-specific binding and incubate for 15 min on ice. Do not wash.</li>
	<li>Add the following antibodies and the respective isotype controls: pacific blue anti-mouse CD45 or rat IgG2a, k (0.25 &#181;g/106 cells in 100 &#181;L volume) and allophycocyanin (APC) anti-mouse EpCAM or rat IgG2b, k (0.5 &#181;g/106 cells in 100 &#181;L volume) monoclonal antibodies. Incubate for 45 min on ice protected from direct light. Add 4.5 mL of cold FACS buffer, mix and spin at 350 x g for 10 min at 4 &#176;C. Discard the FACS buffer and re-suspend the pellet in 300 &#181;L of cold FACS buffer.</li>
	<li>Add propidium iodide (PI) (5 &#181;g/mL) immediately before flow cytometric sorting. 
	NOTE: Brush cells have an irregular shape with several processes that could potentially increase their adherence to filters (Figure 3C). We compared 30 &#956;m to 70 &#956;m and 100 &#956;m filters and found that larger pore filters ensured better yields. In addition, thorough trituration of the cell suspension after papain digestion significantly improves the brush cell yields. We recommend using an 18G needle for initial trituration followed by a smaller bore (21 or 23 G needle) for finer dissociation of cells.</li>
</ol>
4. Flow Cytometry Gating Strategy
<ol>
	<li>Identify cells from debris by forward and side scatter angle. Exclude the doublets using forward scatter height and width and side scatter height and width. The doublets would be the cells that have high width values.</li>
	<li>Within the single cells, identify the live cells as the population that is PI negative.</li>
	<li>Within the live single cells, identify the CD45 low to negative cells based on the isotype control.</li>
	<li>Within the CD45 low/negative cells, identify the EpCAM positive cells that are also eGFP positive (in the FITC channel). This is the population of brush cells (Figure 2A).</li>
</ol>

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The authors have nothing to disclose.

We found that a combination of high-dose dispase treatment for 40 min followed by a short papain treatment (30 min) provides an optimal protocol for tracheal digestion and brush cell isolation. This combination both avoids extensive digestion and produces the highest yield of brush cells, compared to alternate protocols.
While lung digestion to extract hematopoietic cells has classically relied on mild digestive enzymes like collagenase IV15, isolation of epithelial cel...

Brush cells belong to a class of chemosensory epithelial cells characterized by the expression of bitter taste receptors and the taste receptor transduction machinery found in taste bud cells. Unlike taste bud cells, chemosensory epithelial cells are scattered in epithelial surfaces and are referred to as solitary chemosensory cells (SCCs) and microvillous cells in the nasal epithelium1,2, brush cells in the trachea3,4, and tuft cells in the intestine5,6. Epithelial cells expressing bitter taste receptors and the bitter taste transduction machinery are also found in the urethra7,8 and the auditory tube9. Airway brush cells have unique functions in neurogenic and immune airway responses. They are acetylcholine-producing chemosensory cells that evoke protective respiratory reflexes upon activation with bitter compounds and bacterial metabolites like quorum-sensing substances10. Airway brush cells are also the dominant airway epithelial source of IL-25, which regulates aeroallergen-elicited type 2 inflammation in the airways3.
Characterization of the full transcriptome of lower airway brush cells and their response to environmental stimuli has been limited by their low numbers in the tracheal epithelium and very limited numbers beyond the large bronchi10. Techniques used for the isolation of chemosensory cells from the intestinal epithelium have not yielded proportionally high numbers from the trachea, possibly because of the intimate contacts of tracheal brush cells with nerve endings10 or other tissue-specific factors in the respiratory mucosa such as the composition of adherens and tight junction proteins. Recent reports of successful isolation of tracheal brush cells in higher numbers for single cell RNA sequencing analysis employed either a 2 h incubation with papain or an 18 h incubation with pronase11,12. Since longer incubations with digestive enzymes can decrease cell viability and alter the transcriptional profile of cells from digested tissues13, this could bias comparative analysis with other chemosensory epithelial populations.
Here, we report a method for the isolation of tracheal brush cells for RNA sequencing3. Treatment of the trachea with high-dose dispase separates the epithelium from the submucosa. Subsequent digestion of the epithelial sheet with papain allows for excellent recovery of this structural cell.

<table><tbody><tr><td>&lt;strong&gt;Antibodies&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Anti-GFP (Polyclonal goat Ig)</td><td>Abcam</td><td>cat# ab5450</td><td></td></tr><tr><td>APC anti-mouse CD326 (EpCAM)&amp;nbsp; (G8.8)</td><td>Biolegend</td><td>cat#118214</td><td></td></tr><tr><td>APC Rat IgG2a, k isotype control</td><td>Biolegend</td><td>cat#400511</td><td></td></tr><tr><td>DAPI</td><td>Biolegend</td><td>cat#422801</td><td></td></tr><tr><td>Donkey anti-goat IgG (H+L) Highly Cross-Adsorbed Secondary Antibody, Alexa Fluor 488</td><td>Life Technologies/Molecular Probes</td><td>cat#A-11055</td><td></td></tr><tr><td>Normal Goat IgG</td><td>R&amp;amp;D Systems</td><td>cat#AB-108-C</td><td></td></tr><tr><td>Pacific Blue anti-mouse CD45 (30F-11)</td><td>Biolegend</td><td>cat#103126</td><td></td></tr><tr><td>Pacific Blue Rat IgG2b, k isotype control</td><td>Biolegend</td><td>cat#400627</td><td></td></tr><tr><td>TruStain FcX (anti-mouse CD16/32) Antibody</td><td>Biolegend</td><td>cat#101320</td><td></td></tr><tr><td>Chemicals, Peptides, and Recombinant Proteins</td><td></td><td></td><td></td></tr><tr><td>Dispase</td><td>Gibco</td><td>cat# 17105041&amp;nbsp;</td><td></td></tr><tr><td>DNase I</td><td>Sigma&amp;nbsp;</td><td>cat# 10104159001</td><td></td></tr><tr><td>HEPES-Tyrode&amp;rsquo;s Buffer Without Calcium (10 mM HEPES, 135 mM NaCl, 2.8 mM KCl, 1 mM MgCl&lt;sub&gt;2&lt;/sub&gt;, 12 mM NaHCO&lt;sub&gt;3&lt;/sub&gt;, 0.4 mM NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;, 0.25% BSA, 5.5 mM Glucose. Prepared in 18.2 megohms water and filtered through 0.22 &amp;micro;m filter</td><td>Boston BioProducts</td><td>cat# PY-912</td><td></td></tr><tr><td>Tyrode&amp;rsquo;s Solution (HEPES-Buffered) 140 mM NaCl, 5 mM KCl, 25 mM HEPES, 2 mM CaCl&lt;sub&gt;2&lt;/sub&gt;, 2 mM MgCl&lt;sub&gt;2&lt;/sub&gt; and 10 mM glucose. Prepared in 18.2 megohms water and filtered through 0.22 &amp;micro;m filter. )</td><td>Boston BioProducts</td><td>cat# BSS-355</td><td></td></tr><tr><td>L-Cysteine</td><td>Sigma</td><td>cat# C7352</td><td></td></tr><tr><td>Leupeptin trifluoroacetate salt</td><td>Sigma</td><td>cat# L2023</td><td></td></tr><tr><td>Papain from papaya latex</td><td>Sigma</td><td>cat# P3125</td><td></td></tr><tr><td>Propidium iodide&amp;nbsp;</td><td>Sigma</td><td>cat# P4170</td><td></td></tr><tr><td>&lt;strong&gt;Experimental Models: Organisms/Strains&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>ChAT&lt;sup&gt;BAC&lt;/sup&gt;-eGFP (B6.Cg-Tg(RP23-268L19-EGFP)2Mik/J)</td><td>The Jackson Laboratory</td><td>7902</td><td></td></tr><tr><td>&lt;strong&gt;Equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>LSM 800 with Airyscan confocal system&amp;nbsp;on a&amp;nbsp;Zeiss Axio Observer Z1 Inverted Microscope</td><td>Zeiss</td><td></td><td></td></tr><tr><td>LSRFortessa</td><td>BD</td><td>647465</td><td></td></tr><tr><td>&lt;strong&gt;Disposable equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>1.5 mL sterile tubes</td><td>Thomas Scientific</td><td>1157C86</td><td></td></tr><tr><td>5 mL Poysterene Round-bottom Tube, 12 mm x 75 mm style</td><td>Falcon</td><td>14-959-1A</td><td></td></tr><tr><td>50 mL Polypropylene conical tube, 30 mm x 115 mm style</td><td>Falcon</td><td>352098</td><td></td></tr><tr><td>Feather Disposable Scalpel no.12</td><td>Fisher Scientific</td><td>NC9999403</td><td></td></tr><tr><td>Petri dish, 100 mm x 15 mm Style</td><td>Falcon</td><td>351029</td><td></td></tr><tr><td>Sterile cell strainer, 100 &amp;mu;m</td><td>Fisherbrand</td><td>cat#22363549</td><td></td></tr></tbody></table>

isolation and quantitative evaluation of brush cells from mouse tracheas

Tracheal brush cells are cholinergic chemosensory epithelial cells poised to transmit signals from the airway lumen to the immune and nervous systems. They are part of a family of chemosensory epithelial cells which include tuft cells in the intestinal mucosa, brush cells in the trachea, and solitary chemosensory and microvillous cells in the nasal mucosa. Chemosensory cells in different epithelial compartments share key intracellular markers and a core transcriptional signature, but also display significant transcriptional heterogeneity, likely reflective of the local tissue environment. Isolation of tracheal brush cells from single cell suspensions is required to define the function of these rare epithelial cells in detail, but their isolation is challenging, potentially due to the close interaction between tracheal brush cells and nerve endings or due to airway-specific composition of tight and adherens junctions. Here, we describe a procedure for isolation of brush cells from mouse tracheal epithelium. The method is based on an initial separation of tracheal epithelium from the submucosa, allowing for a subsequent shorter incubation of the epithelial sheet with papain. This procedure offers a rapid and convenient solution for flow cytometric sorting and functional analysis of viable tracheal brush cells.

This procedure has been successfully implemented to isolate tracheal brush cells for RNA sequencing3. After isolation of the trachea and digestion of the tissue with a 2-step protocol (Figure 1), cells were collected and stained with fluorescently-labeled CD45 and EpCAM after exclusion of dead cells with PI. After gating out doublets based on forward and side scatter characteristics, we defined brush cells as low/negative for CD45, positive for EpCAM and positive for ...

Watch this Scientific Journal Video about Isolation and Quantitative Evaluation of Brush Cells from Mouse Tracheas at JoVE.com

Isolation and Quantitative Evaluation of Brush Cells from Mouse Tracheas

Brush cells are rare cholinergic chemosensory epithelial cells found in the na&#239;ve mouse trachea. Due to their limited numbers, ex vivo evaluation of their functional role in airway immunity and remodeling is challenging. We describe a method for isolation of tracheal brush cells by flow cytometry.

Tracheal brush cells are cholinergic chemosensory epithelial cells poised to transmit signals from the airway lumen to the immune and nervous systems. ...

isolation-quantitative-evaluation-brush-cells-from-mouse

Research

JoVE Journal

Immunology and Infection

7.2K Views.  Harvard Medical School. We ontwikkelden een procedure voor isolatie van tracheale borstelcellen, zeldzame gespecialiseerde chemosensorische epitheelcellen van choline acetyltransferase fluorescerende reporter muizen. Deze methode is gebaseerd op een eerste scheiding van tracheale epitheel van de submucosa waardoor een latere kortere incubatie van het epitheelblad met papaïne mogelijk is. Dit zorgt voor een robuust herstel van tracheale borstelcellen en is met succes gebruikt voor isolatie en transcriptie analyse va...

Video: Isolation and Quantitative Evaluation of Brush Cells from Mouse Tracheas

Isolation of Mouse Respiratory Epithelial Cells and Exposure to Experimental Cigarette Smoke at Air Liquid Interface  

Pulmonary epithelial cells can be isolated from the respiratory tract of mice and cultured at air-liquid interface (ALI) as a model of differentiated respiratory epithelium. A protocol is described for isolating and exposing these cells to mainstream cigarette smoke (CS), in order to study epithelial cell responses to CS exposure. The protocol consists of three parts: the isolation of airway epithelial cells from mouse trachea, the culturing of these cells at air-liquid interface (ALI) as fully differentiated epithelial cells, and the delivery of calibrated mainstream CS to these cells in culture. The ALI culture system allows the culture of respiratory epithelia under conditions that more closely resemble their physiological setting than ordinary liquid culture systems. The study of molecular and lung cellular responses to CS exposure is a critical component of understanding the impact of environmental air pollution on human health. Research findings in this area may ultimately contribute towards understanding the etiology of chronic obstructive pulmonary disease (COPD), and other tobacco-related diseases, which represent major global health problems.

Isolation of Mouse Respiratory Epithelial Cells and Exposure to Experimental Cigarette Smoke at Air Liquid Interface

Pulmonary epithelial cells can be isolated from the respiratory tract of mice and cultured at air-liquid interface as a model of differentiated respiratory epithelium. A protocol is described for isolating, culturing and exposing these cells to mainstream cigarette smoke, in order to study molecular responses to this environmental toxin.

Pulmonary epithelial cells can be isolated from the respiratory tract of mice and cultured at air-liquid interface (ALI) as a model of differentiated ...

Medicine

In vitro Measurements of Tracheal Constriction Using Mice

Transgenic and knockout mice have been powerful tools for the investigation of the physiology and pathophysiology of airways1,2. In vitro tensometry of isolated tracheal preparations has proven to be a useful assay of airway smooth muscle (ASM) contractile response in genetically modified mice. These in vitro tracheal preparations are relatively simple, provide a robust response, and retain both functional cholinergic nerve endings and muscle responses, even after long incubations.
Tracheal tensometry also provides a functional assay to study a variety of second messenger signaling pathways that affect contraction of smooth muscle. Contraction in trachea is primarily mediated by parasympathetic, cholinergic nerves that release acetylcholine onto ASM (Figure 1). The major ASM acetylcholine receptors are muscarinic M2 and M3 which are Gi/o and Gq coupled receptors, respectively3,4,5. M3 receptors evoke contraction by coupling to Gq to activate phospholipase C, increase IP3 production and IP3-mediated calcium release from the sarcoplasmic reticulum3,6,7. M2/Gi/o signaling is believed to enhance contractions by inhibition of adenylate cyclase leading to a decrease in cAMP levels5,8,9,10. These pathways constitute the so called "pharmaco-contraction coupling" of airway smooth muscle11. In addition, cholinergic signaling through M2 receptors (and modulated by M3 signaling) involves pathways that depolarize the ASM which in turn activate L-type, voltage-dependent calcium channels (Figure 1) and calcium influx (so called "excitation-contraction coupling")4,7. More detailed reviews on signaling pathways controlling airway constriction can be found4,12. The above pathways appear to be conserved between mice and other species. However, mouse tracheas differ from other species in some signaling pathways. Most prominent is their lack of contractile response to histamine and adenosine13,14, both well-known ASM modulators in humans and other species5,15.
Here we present protocols for the isolation of murine tracheal rings and the in vitro measurement of their contractile output. Included are descriptions of the equipment configuration, trachea ring isolation and contractile measurements. Examples are given for evoking contractions indirectly using high potassium stimulation of nerves and directly by depolarization of ASM muscle to activate voltage-dependent calcium influx (1. high K+, Figure 1). In addition, methods are presented for stimulations of nerves alone using electric field stimulation (2. EFS, Figure 1), or for direct stimulation of ASM muscle using exogenous neurotransmitter applied to the bath (3. exogenous ACH, Figure 1). This flexibility and ease of preparation renders the isolated trachea ring model a robust and functional assay for a number of signaling cascades involved in airway smooth muscle contraction.

In vitro Measurements of Tracheal Constriction Using Mice

Transgenic mice have been extremely useful in ascribing physiological function to genes. As such, research in general, and functional studies of airway, in particular, have undergone a remarkable shift toward murine models. Here we provide protocols for in vitro trachea constriction studies to evaluate smooth muscle function in murine airway.

Transgenic and knockout mice have been powerful tools for the investigation of the physiology and pathophysiology of airways1,2. In vitro tensometry of ...

Flow Cytometric Isolation of Primary Murine Type II Alveolar Epithelial Cells for Functional and Molecular Studies

Throughout the last years, the contribution of alveolar type II epithelial cells (AECII) to various aspects of immune regulation in the lung has been increasingly recognized. AECII have been shown to participate in cytokine production in inflamed airways and to even act as antigen-presenting cells in both infection and T-cell mediated autoimmunity 1-8. Therefore, they are especially interesting also in clinical contexts such as airway hyper-reactivity to foreign and self-antigens as well as infections that directly or indirectly target AECII. However, our understanding of the detailed immunologic functions served by alveolar type II epithelial cells in the healthy lung as well as in inflammation remains fragmentary. Many studies regarding AECII function are performed using mouse or human alveolar epithelial cell lines 9-12. Working with cell lines certainly offers a range of benefits, such as the availability of large numbers of cells for extensive analyses. However, we believe the use of primary murine AECII allows a better understanding of the role of this cell type in complex processes like infection or autoimmune inflammation. Primary murine AECII can be isolated directly from animals suffering from such respiratory conditions, meaning they have been subject to all additional extrinsic factors playing a role in the analyzed setting. As an example, viable AECII can be isolated from mice intranasally infected with influenza A virus, which primarily targets these cells for replication 13. Importantly, through ex vivo infection of AECII isolated from healthy mice, studies of the cellular responses mounted upon infection can be further extended.
Our protocol for the isolation of primary murine AECII is based on enzymatic digestion of the mouse lung followed by labeling of the resulting cell suspension with antibodies specific for CD11c, CD11b, F4/80, CD19, CD45 and CD16/CD32. Granular AECII are then identified as the unlabeled and sideward scatter high (SSChigh) cell population and are separated by fluorescence activated cell sorting 3.
In comparison to alternative methods of isolating primary epithelial cells from mouse lungs, our protocol for flow cytometric isolation of AECII by negative selection yields untouched, highly viable and pure AECII in relatively short time. Additionally, and in contrast to conventional methods of isolation by panning and depletion of lymphocytes via binding of antibody-coupled magnetic beads 14, 15, flow cytometric cell-sorting allows discrimination by means of cell size and granularity. Given that instrumentation for flow cytometric cell sorting is available, the described procedure can be applied at relatively low costs. Next to standard antibodies and enzymes for lung disintegration, no additional reagents such as magnetic beads are required. The isolated cells are suitable for a wide range of functional and molecular studies, which include in vitro culture and T-cell stimulation assays as well as transcriptome, proteome or secretome analyses 3, 4.

We describe the rapid isolation of primary murine type II alveolar epithelial cells (AECII) by flow cytometric negative selection. These AECII show high viability and purity and are suitable for a wide range of functional and molecular studies regarding their role in respiratory conditions such as autoimmune or infectious diseases.

Throughout  the last years, the contribution of alveolar type II epithelial cells (AECII)  to various aspects of immune regulation in the lung has been ...

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 ...

An Improved Method for Rapid Intubation of the Trachea in Mice

Despite some anatomical and physiological differences, mouse models continue to be an essential tool for studying human lung disease. Bleomycin toxicity is a commonly used model to study both acute lung injury and fibrosis, and multiple methods have been developed for administering bleomycin (and other toxic agents) into the lungs. However, many of these approaches, such as transtracheal instillation, have inherent drawbacks, including the need for strong anesthetics and survival surgery. This paper reports a quick, reproducible method of intratracheal intubation that involves mild inhaled anesthesia, visualization of the trachea, and the use of a surrogate spirometer to confirm exposure. As a proof of concept, 8-12 week old C57BL/6 mice were administered either 2.0 U/kg of bleomycin or an equivalent volume of PBS, and both damage and fibrotic endpoints were measured post-exposure. This procedure allows researchers to treat a large cohort of mice in a relatively short period with little expense and minimal post-procedure care.

This article presents a rapid and simple method for administering bleomycin directly into the mouse trachea via intubation. Key advantages of this method are that it is highly reproducible, easy to master, and does not require specialized equipment or lengthy recovery times.

Despite some anatomical and physiological differences, mouse models continue to be an essential tool for studying human lung disease. Bleomycin toxicity ...

Design of a Biocompatible Drug-Eluting Tracheal Stent in Mice with Laryngotracheal Stenosis

Laryngotracheal stenosis (LTS) is a pathologic narrowing of the subglottis and trachea leading to extrathoracic obstruction and significant shortness of breath. LTS results from mucosal injury from a foreign body in the trachea, leading to tissue damage and a local inflammatory response that goes awry, leading to the deposition of pathologic scar tissue. Treatment for LTS is surgical due to the lack of effective medical therapies. The purpose of this method is to construct a biocompatible stent that can be miniaturized to place into mice with LTS. We demonstrated that a PLLA-PCL (70% poly-L-lactide and 30% polycaprolactone) construct had optimal biomechanical strength, was biocompatible, practicable for an in vivo placement stent, and capable of eluting drug. This method provides a drug delivery system for testing various immunomodulatory agents to locally inhibit inflammation and reduce airway fibrosis. Manufacturing the stents takes 28&#8722;30 h and can be reproduced easily, allowing for experiments with large cohorts. Here we incorporated the drug rapamycin within the stent to test its effectiveness in reducing fibrosis and collagen deposition. Results revealed that PLLA-PCL tents showed reliable rapamycin release, were mechanically stable in physiological conditions, and were biocompatible, inducing little inflammatory response in the trachea. Further, the rapamycin-eluting PLLA-PCL stents reduced scar formation in the trachea in vivo.

Laryngotracheal stenosis results from pathologic scar deposition that critically narrows the tracheal airway and lacks effective medical therapies. Using a PLLA-PCL (70% poly-L-lactide and 30% polycaprolactone) stent as a local drug delivery system, potential therapies aimed at decreasing scar proliferation in the trachea can be studied.

Laryngotracheal stenosis (LTS) is a pathologic narrowing of the subglottis and trachea leading to extrathoracic obstruction and significant shortness of ...

Bioengineering

Isolation of Basal Cells and Submucosal Gland Duct Cells from Mouse Trachea

The large airways are directly in contact with the environment and therefore susceptible to injury from toxins and infectious agents that we breath in 1. The large airways therefore require an efficient repair mechanism to protect our bodies. This repair process occurs from stem cells in the airways and isolating these stem cells from the airways is important for understanding the mechanisms of repair and regeneration. It is also important for understanding abnormal repair that can lead to airway diseases 2. The goal of this method is to isolate a novel stem cell population from the mouse tracheal submucosal gland ducts and to place these cells in in vitro and in vivo model systems to identify the mechanisms of repair and regeneration of the submucosal glands 3. This production shows methods that can be used to isolate and assay the duct and basal stem cells from the large airways 3.This will allow us to study diseases of the airway, such as cystic fibrosis, asthma and chronic obstructive pulmonary disease. Currently, there are no methods for isolation of submucosal gland duct cells and there are no in vivo models to study the regeneration of submucosal glands.

Here we demonstrate our protocol for isolation of basal and submucosal gland duct cells from mouse tracheas. We also demonstrate the method of injecting stem cells into the dorsal mouse fat pad to create an in vivo model of submucosal gland regeneration.

The large airways are directly in contact with the environment and therefore susceptible to injury from toxins and infectious agents that we breath in 1. ...

Biology

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  ...

Isolating Bronchial Epithelial Cells from Resected Lung Tissue for Biobanking and Establishing Well-Differentiated Air-Liquid Interface Cultures

The airway epithelial cell layer forms the first barrier between lung tissue and the outside environment and is thereby constantly exposed to inhaled substances, including infectious agents and air pollutants. The airway epithelial layer plays a central role in a large variety of acute and chronic lung diseases, and various treatments targeting this epithelium are administered by inhalation. Understanding the role of epithelium in pathogenesis and how it can be targeted for therapy requires robust and representative models. In vitro epithelial culture models are increasingly being used and offer the advantage of performing experiments in a controlled environment, exposing the cells to different kinds of stimuli, toxicants, or infectious agents. The use of primary cells instead of immortalized or tumor cell lines has the advantage that these cells differentiate in culture to a pseudostratified polarized epithelial cell layer with a better representation of the epithelium compared to cell lines.
Presented here is a robust protocol, that has been optimized over the past decades, for the isolation and culture of airway epithelial cells from lung tissue. This procedure allows successful isolation, expansion, culture, and mucociliary differentiation of primary bronchial epithelial cells (PBECs) by culturing at the air-liquid interface (ALI) and includes a protocol for biobanking. Furthermore, the characterization of these cultures using cell-specific marker genes is described. These ALI-PBEC cultures can be used for a range of applications, including exposure to whole cigarette smoke or inflammatory mediators, and co-culture/infection with viruses or bacteria. 
The protocol provided in this manuscript, illustrating the procedure in a step-by-step manner, is expected to provide a basis and/or reference for those interested in implementing or adapting such culture systems in their laboratory.

Presented here is a reproducible, affordable, and robust method for the isolation and expansion of primary bronchial epithelial cells for long-term biobanking and the generation of differentiated epithelial cells by culture at the air-liquid interface.

The airway epithelial cell layer forms the first barrier between lung tissue and the outside environment and is thereby constantly exposed to inhaled ...

Noninvasive Real-Time Whole-Body Plethysmography for Cough Sensitivity in Mice

Methods for Detecting Cough and Airway Inflammation in Mice

Chronic cough, which lasts for more than 8 weeks, is one of the most common complaints requiring medical attention, and patients suffer from a huge socioeconomic burden and a marked decrement in quality of life. Animal models can mimic the complex pathophysiology of the cough and are important tools for cough research. The detection of cough sensitivity and airway inflammation is of great significance for studying the complex pathological mechanism of cough. This article describes the measurement of cough using a noninvasive and real-time whole-body plethysmography (WBP) system and the normative procedures for harvesting tissue samples (including blood, lung, spleen, and trachea) of mice. It introduces some methods to assess airway inflammation, including pathological changes in hematoxylin and eosin (HE)-stained lung and trachea sections, the total protein concentration, the uric acid concentration, and the lactate dehydrogenase (LDH) activity in the supernatant of bronchoalveolar lavage fluid (BALF), and the leukocytes and differential cell counts of BALF. These methods are reproducible and serve as valuable tools to study the complex pathophysiology of cough.

Here, we describe the measurement of cough using a noninvasive and real-time whole-body plethysmography (WBP) system and the normative procedures for harvesting tissue samples of mice and introduce some methods to assess airway inflammation.

Chronic cough, which lasts for more than 8 weeks, is one of the most common complaints requiring medical attention, and patients suffer from a huge ...

Isolation and Quantitative Evaluation of Brush Cells from Mouse Tracheas

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