Name: characterization of immune cells and proinflammatory mediators in the pulmonary environment
Uploaded: 2020-06-24T12:30:00.000+00:00
Duration: 9 min
Description: Watch this Scientific Journal Video about Characterization of Immune Cells and Proinflammatory Mediators in the Pulmonary Environment at JoVE.com

This work was supported by NIH grant P20 GM109098 and the Innovation Award Program from Praespero to Edwin Wan. Flow Cytometry experiments were performed in the WVU Flow Cytometry &#38; Single Cell Core Facility, which is&#160;supported by NIH grants S10 OD016165, U57 GM104942, P30 GM103488, and P20 GM103434.

All protocols and procedures performed were approved by the Institutional Animal Care and Use Committee (IACUC) of West Virginia University. The mice were housed under specific-pathogen-free conditions in the vivarium at West Virginia University. &#160;
1. Preparation of solutions
<ol>
	<li>Prepare perfusion buffer (phosphate buffered saline, PBS). Use approximately two 10 mL aliquots of ice-cold PBS per mouse.</li>
	<li>Prepare lung cell medium/FACS buffer. FACS buffer contains PBS supplemented with 1% fetal bovine serum (FBS). Keep the medium cold for the entire process of lung excision and transfer. Prepare approximately 8 mL per lung sample. Prepare fresh medium/FACS buffer prior to the experiments.</li>
	<li>Prepare dissociation buffer for single cell isolation. Buffer contains 1 mg/mL collagenase D and 200 &#181;g/mL DNase I in Hank&#8217;s buffered salt solution (HBSS). Prepare fresh from the stock solution (100 mg/mL collagenase D and 10 mg/mL DNase I) prior to the experiments. Approximately 6 mL per lung sample is needed. Ensure buffer reaches room temperature prior to the use.</li>
	<li>Prepare homogenization buffer for lung tissue homogenization. Buffer contains PBS and 1x proteinase and phosphatase inhibitor cocktail (stock = 100x). Freshly prepare prior to the experiments. Keep the buffer cold during the entire homogenization process. Approximately 200 &#181;L of the buffer is sufficient for homogenizing a single right lobe of the lungs.</li>
</ol>
2. Transient middle cerebral artery occlusion (tMCAO)
NOTE: Procedures for tMCAO via monofilament insertion to the middle cerebral artery were documented in detail previously10. In these experiments, 8- to 12-week-old male C57BL/6J mice, weighing 25-30 g, were used.
<ol>
	<li>In brief, subcutaneously administer 2 mg/kg meloxicam to the mice before surgery. Deeply anesthetize the mice in an induction chamber using 5% isoflurane. Confirm deep anesthetization using the toe-pinch method. Maintain anesthesia with 1-2% isoflurane during the surgery using a nose cone. Maintain the body temperature at 36.5 - 38 &#176;C throughout the procedure by placing a warm blanket underneath the mice. All surgical tools should be autoclave-sterilized a day prior to the surgery.</li>
	<li>Shave and prepare the skin on the ventral neck using 70% ethanol followed by chlorhexidine scrub. Subcutaneously administer 2 mg/kg bupivacaine over the incision area prior to making the first incision. Use eye ointment to cover the eyes to prevent dryness. Make a midline neck incision. Gently pull aside soft tissues around the trachea.</li>
	<li>Identify the left common carotid artery and the external carotid artery.</li>
	<li>Apply a temporary suture (size 6/0) to the common carotid artery to stem the flow of the blood. Make an incision in the external carotid artery.</li>
	<li>Insert a silicone rubber-coated monofilament (size 6-0) into the external carotid artery, then advance the monofilament to the middle cerebral artery. Leave the monofilament in the middle cerebral artery for 60 min (occlusion).</li>
	<li>Monitor the rate of blood flow using Laser Doppler Flowmetry. A reduction in the flow rate to the middle cerebral artery that is &#62; 80% indicates a successful occlusion.</li>
	<li>Following the 60 min of occlusion, remove the monofilament to allow for the reperfusion to occur. Close the incision.</li>
	<li>In sham-operated mice, perform steps 2.1-2.6 without the insertion of the monofilament. In these mice, the rate of blood flow should remain steady during the entire procedure.</li>
	<li>Monitor the mice daily and measure the neurological deficits using standard scoring criteria11. 0 &#8211; no neurological deficit; 1 &#8211; retracts contralateral forepaw when lifted by the tail; 2- circles to the contralateral when lifted by the tail; 3- falling to the contralateral while walking; 4 &#8211; does not walk spontaneously or is comatose; 5 &#8211; dead.</li>
	<li>Provide subcutaneous injections of normal saline and analgesic (meloxicam, 2 mg/kg) every 24 hours for 3 days following the surgery.</li>
	<li>Isolate the lungs 24 and 72 h following tMCAO for the downstream analysis.</li>
</ol>
3. Harvesting the lung tissues
<ol>
	<li>Deeply anesthetize the mouse by intraperitoneal (i.p.) injection of 100 mg/kg of ketamine and 10 mg/kg of xylazine. Confirm deep anesthetization using the toe-pinch method.</li>
	<li>To perform the whole body transcardial perfusion12, pin the mouse to the surgical platform and wet fur with 70% ethanol to reduce fur distribution into the body cavity. Use fine dissection scissors to make a transverse incision in the caudal abdomen and then a midline incision of the peritoneum, stopping at the thoracic cavity prior to piercing the diaphragm.
	<ol>
		<li>Do not cut through the muscle/hide into the peritoneal cavity and do not continue incision past abdomen.</li>
	</ol>
	</li>
	<li>Stop the incision as soon as the thoracic cavity has been reached before puncturing the diaphragm. Take care not to damage the heart and lungs which may compromise perfusion quality or cell recovery</li>
	<li>Using fine dissection scissors make a midline incision of the peritoneum stopping at the thoracic cavity prior to piercing the diaphragm.</li>
	<li>Grasp the mouse by the distal end of the sternum using forceps and lift upward while cutting through the diaphragm being certain not to damage the heart, lungs, or vasculature. Once organs are visible make an incision through the rib cage and separate and pin back the rib case so that the heart and lungs are fully exposed.</li>
	<li>Carefully make a small incision in the right atrium near the aortic arch. This will serve as the outflow for the perfusion.</li>
	<li>Immediately after, gently insert a 25 G x 5/8 needle attached to a syringe filled with 10 mL of cold PBS into the distal superior surface of the left ventricle. Use care not to insert through the ventricular septum into the right ventricle. Needle should be barely inserted into the left ventricle. 
	NOTE: If ventricular septum is pierced, fluid will rush out from the mouse&#8217;s nose. This will decrease the perfusion quality and result in cell loss.</li>
	<li>If desired, a clamp may be used to hold the needle securely in place in the left ventricle during the perfusion.</li>
	<li>Steadily but gently push 10 mL of cold PBS into the heart. Mouse tissue should begin to perfuse and clear off the blood as fluid exits from the right atrium.
	<ol>
		<li>Push a second aliquot of 10 mL of PBS until sufficient blood clearance has occurred. Liver will have a light brown appearance and lungs will transition from a reddish pink to mostly white in color.</li>
	</ol>
	</li>
	<li>Using forceps and fine dissection scissors, carefully remove all surrounding tissues including the heart, trachea, esophagus, thymus, connective tissue, lymph nodes, and large bronchials.</li>
	<li>Separate individual lung lobes and place the lobes into a Petri dish on ice containing a 1-2 mL aliquot of cold lung cell medium.</li>
</ol>
4. Homogenization of lung tissue for multiplex bead arrays using bead homogenizer
<ol>
	<li>Pre-chill a 2 mL conical screw cap tube containing 3 sterile 2.3 mm zirconia/silica beads. Add 200 &#181;L of cold homogenization buffer to the tube.</li>
	<li>Weigh the lobe, transfer the lobe to the pre-chilled conical tube containing the beads and homogenization buffer. 
	NOTE: Approximately 50-100 mg of tissue homogenized in 200 &#181;L of buffer will be sufficient for detecting cytokine and chemokine expression in the sample.
	<ol>
		<li>Homogenize the lobe using a bead-based homogenizer (see Table of Materials) at 4,000 rpm for 2 min.</li>
		<li>Ensure the tissue is completely homogenized. Increase the time if needed.</li>
	</ol>
	</li>
	<li>Following homogenization, centrifuge the tube in a pre-chilled (4 &#176;C) micro-centrifuge at 15,870 x g for 3 min.</li>
	<li>Transfer the supernatants to a pre-chilled 1.5 mL microtube.</li>
	<li>Place the sample directly on ice for immediate use or store the sample at -80 &#176;C for future use. Avoid multiple freeze-thaw cycles.</li>
</ol>
5. Lung tissue dissociation and single cell isolation
<ol>
	<li>Pour the lung cell medium off, of the remaining lung lobes to avoid dilution of the dissociation buffer. Then, carefully insert a 1 mL syringe with 25G and 5/8 needle containing 1 mL of the dissociation buffer into lung tissue. &#160;</li>
	<li>Inject small fractions (approximately 1/4th to 1/5th) of the dissociation buffer into each lung lobe and gently inflate. 
	NOTE: Most of the buffer will rush out soon after inflating as the lobe has been excised.</li>
	<li>Repeat the injection of dissociation buffer 1-2x.</li>
	<li>Incubate the lung lobes with dissociation buffer for 2 min at room temperature.</li>
	<li>Finely mince the lung lobes into small pieces using fine dissection scissors.</li>
	<li>Transfer the minced lung pieces with the dissociation buffer to a 15 mL centrifuge tube. &#160;Supplement with additional dissociation buffer to a total of 6 mL. Vortex vigorously for 1 min.</li>
	<li>Incubate the sample for 45 min at 37 &#176;C. Vortex vigorously every 7-8 min.</li>
	<li>After 45 min of incubation, vortex the sample vigorously for 1 min for optimal results.</li>
	<li>Further dissociate the sample by passing through a 100 &#181;m cell strainer to remove residual, undesired connective, and interstitial tissue.</li>
	<li>Centrifuge the sample for 10 min at 380 x g, discard the supernatant.</li>
	<li>Add 5 mL of cold lung cell medium to the sample. Resuspend the cell pellet by gentle vortexing.</li>
	<li>Centrifuge the sample for 10 min at 380 x g, discard the supernatant.</li>
	<li>Add 1 mL of cold lung cell medium. Resuspend the cell pellet by gentle vortexing.</li>
	<li>Pass through a 100 &#181;m cell strainer, and count cells.</li>
</ol>
6. Flow cytometric analysis for the lung immune cell niche
<ol>
	<li>Seed 1-2 x 106 cells/antibody set into a 96 well round bottom plate (3 sets in total). Centrifuge the cells for 3 min at 830 x g. Discard supernatants.</li>
	<li>Resuspend the cell pellet in 50 &#181;L of cold PBS containing fixable live/dead stain. Prepare the stain according to the manufacturer&#8217;s instructions.</li>
	<li>Incubate the cells at 4 &#176;C for 20 min protected from light. Centrifuge the cells for 3 min at 830 x g. Discard supernatants.&#160;&#160;</li>
	<li>Resuspend the cell pellet with 25 &#181;L of cold FACS buffer containing 5 &#181;g/mL of anti-mouse CD16/32 antibody. Incubate at 4 &#176;C for 10-15 min.</li>
	<li>Add 25 &#181;L of cold FACS buffer containing antibody combinations listed in Table 1 to the cells. Mix by gentle pipetting. 
	NOTE: Total volume of FACS buffer for the antibody staining is 50 &#181;L. Therefore, when preparing a master mix of antibodies in 25 &#181;L, the concentration of antibodies should be double the final concentration. &#160;</li>
	<li>Incubate the cells at 4 &#176;C for 20 min protected from light. Centrifuge the cells for 3 min at 830 x g. Discard supernatants.</li>
	<li>Resuspend the cells with 100 &#181;L of cold FACS buffer. Centrifuge the cells for 3 min at 830 x g. Discard supernatants.</li>
	<li>Resuspend the cells with 100 &#181;L of cold FACS buffer. Transfer to a polystyrene round-bottom 5 mL FACS tube containing 150 &#181;L of additional FACs buffer. Analyze the sample immediately using a flow cytometer. 
	NOTE: Cell fixation using the following steps allows samples to be analyzed later.</li>
	<li>Following step 6.7., resuspend the sample with 100 &#181;L of 1% paraformaldehyde in PBS.</li>
	<li>Incubate the sample at 4 &#176;C for 15 min protected from light. Centrifuge the cells for 3 min at 830 x g. Discard supernatants.</li>
	<li>Resuspend the cells with 100 &#181;L of cold FACS buffer. Centrifuge the cells for 3 min at 830 x g. Discard supernatants.</li>
	<li>Resuspend the cells with 100 &#181;L of cold FACS buffer. Store the cells at 4 &#176;C, protected from light. Analyze the sample within 72 h.</li>
</ol>
7. Multiplex bead arrays for cytokine and chemokine detection
NOTE: Commercially available proinflammatory chemokine and inflammation multiplex panels (see Table of Materials) were used to determine the expression of chemokines and cytokines in the lungs following tMCAO induction according to manufacturer protocol, which has been described in detail13.
<ol>
	<li>In brief, thaw the analytes on ice if they were stored at -80 &#176;C after tissue homogenization. Centrifuge the sample at 590 x g for 2 min at 4 &#176;C before use to remove any residual tissue.</li>
	<li>Generate a standard curve by serially diluting the standard cocktail (C7) which is at a known concentration of 10 ng/mL into 7 standards (C1-6) with the last standard being assay buffer alone (C0).</li>
	<li>Once all reagents have warmed to room temperature, add 25 &#181;L of each standard and 25 &#181;L of assay buffer to a V bottom 96 well plate.&#160;</li>
	<li>To each sample well, add 25 &#181;L of lung homogenate (analyte) and 25 &#181;L of assay buffer to a V bottom 96 well plate</li>
	<li>Vortex the pre-coated beads vigorously, then add 25 &#181;L of pre-coated beads to each of the standard and sample wells.</li>
	<li>Seal the plate and protect from light by covering with foil. Shake the plate at 110 rpm for 2 h at room temperature.</li>
	<li>Centrifuge at 230 x g for 5 min. Add 200 &#181;L of wash buffer at 1x (stock solution is 20x, dilute to 1x with water). Incubate for 1 min.</li>
	<li>Centrifuge at 230 x g for 5 min. Resuspend the standard and sample with 25 &#181;L detection antibodies.</li>
	<li>Seal and cover the plate to protect from light. Shake plate at 110 rpm for 1 h at room temperature.</li>
	<li>Add 25 &#181;L of streptavidin-phycoerythrin (SA-PE) to each standard and sample well.</li>
	<li>Seal and cover the plate to protect from light. Shake at 110 rpm for 30 min at room temperature.</li>
	<li>Spin the plate at 230 x g for 5 min. Resuspend the standard and samples in 1x wash buffer.</li>
	<li>Transfer to a well-labeled polystyrene round-bottom 5 mL FACS tube containing an additional 150 &#181;L of 1x wash buffer.</li>
	<li>Determine the PE signal fluorescence intensity for the standards and samples using a flow cytometer.</li>
	<li>Construct a standard curve for each chemokine and cytokine using the mean fluorescence intensity (MFI) of PE against the pre-determined concentrations.</li>
	<li>Determine the concentration of each chemokine and cytokine using the standard curve and the MFI from each sample. The multiplex panel provides data analysis software that can be used to determine the concentration of each analyte. The weight of the apical/superior lobe is used to determine the concentration of cytokines or chemokine per milligram of tissue. &#160;&#160;</li>
</ol>

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The authors have nothing to disclose.

The protocols described here allow for the identification of lung immune cell types and the expression of chemokines or cytokines in the same mouse. If a histopathology study is desired, an individual lobe can be removed and fixed for that purpose prior to proceeding to the single cell isolation steps. One limitation of this method is that this approach may not be suitable in some disease settings if the change in the immune cell composition and the expression of chemokines and/or cytokines are anticipated to be unequall...

The lungs are a barrier organ, exposed to the external environment and, therefore, are constantly receiving immunological challenges such as pathogens and allergens1. The activation of lung-resident immune cells and the infiltration of immune cells from the periphery are required to clear pathogens from the pulmonary environment. Additionally, lung-resident immune cells maintain tolerance to commensal bacteria, suggesting that these cells play a role in pathogen clearance and maintaining homeostasis1. Alveolar and interstitial macrophages are among the lung-resident sentinel immune cells that sense pathogens via pattern recognition receptors and clear these pathogens by phagocytosis2. Lung-resident dendritic cells bridge the innate and adaptive immune response through antigen presentation3. In addition, activated local innate immune cells produce cytokines and chemokines that amplify the inflammatory response and stimulate the infiltration of immune cells such as monocytes, neutrophils, and lymphocytes into the lungs1. Ischemic stroke has been shown to modify systemic immunity and lead to increased susceptibility to pulmonary infection; however, few studies have evaluated the pulmonary compartment following ischemic stroke, though some studies have examined it during &#160;inflammatory conditions4,5,6,7,8,9. The goal of the methods described herein is to simultaneously determine lung pathology, immune cell composition, and the levels of cytokine and chemokine expression in the lungs to evaluate alterations to the pulmonary compartment and assess potential alterations to the pulmonary immune response following ischemic attack.
Described here is a protocol for obtaining single cell suspensions from the lungs of the mice to identify 13 types of immune cells. This protocol is based on tissue digestion with collagenase D without the need of an automated tissue dissociator. Additionally, we developed a protocol to prepare tissue homogenates that can be used to determine the expression levels of 13 different cytokines or chemokines using flow cytometry-based multiplex bead arrays. This protocol was successfully used to investigate the effects of ischemic stroke on pulmonary immunity and can be used in other disease models as well.

<table><tbody><tr><td>B220-APC, clone RA3-6B2</td><td>Biolegend</td><td>103212</td><td>1:200 dilution</td></tr><tr><td>Beadbug 3 position bead homogenizer</td><td>Benchmark Scientific</td><td>D1030</td><td>Tissue homogenizer</td></tr><tr><td>CCR2-BV421, clone SA203G11</td><td>Biolegend</td><td>150605</td><td>1:200 dilution</td></tr><tr><td>CD103-BV421, clone 2E7</td><td>Biolegend</td><td>121422</td><td>1:200 dilution</td></tr><tr><td>CD11b-PE/Cy7, clone M1/70</td><td>Biolegend</td><td>101216</td><td>1:400 dilution</td></tr><tr><td>CD11c-PE/Cy7, clone N418</td><td>Biolegend</td><td>117318</td><td>1:200 dilution</td></tr><tr><td>CD11c-Percp/Cy5.5, clone N418</td><td>Biolegend</td><td>117328</td><td>1:200 dilution</td></tr><tr><td>CD4-BV421, clone GK1.5</td><td>Biolegend</td><td>100443</td><td>1:200 dilution</td></tr><tr><td>CD45-FITC, clone 30-F11</td><td>Biolegend</td><td>103108</td><td>1:200 dilution</td></tr><tr><td>CD64-APC, clone X54-5/7.1</td><td>Biolegend</td><td>139306</td><td>1:200 dilution</td></tr><tr><td>CD8-PE, clone 53-6.7</td><td>Biolegend</td><td>100708</td><td>1:800 dilution</td></tr><tr><td>Collagenase D</td><td>Sigma Aldrich</td><td>11088882001</td><td>Component in the dissociation buffer</td></tr><tr><td>Conical screw cap tube</td><td>ThermoFisher</td><td>02-681-344</td><td>Tube for tissue homogenization</td></tr><tr><td>DNase I</td><td>Sigma Aldrich</td><td>10104159001</td><td>Component in the dissociation buffer</td></tr><tr><td>Fc block CD16/32 antibody</td><td>Biolegend</td><td>101320</td><td>1:100 dilution</td></tr><tr><td>genlteMACS dissociator</td><td>Miltenyi Biotec</td><td>130-093-235</td><td>Comparsion of lung digestion with or without mechanical dissociator</td></tr><tr><td>gentleMACS C tubes</td><td>Miltenyi Biotec</td><td>130-093-237</td><td>Tube for tissue disscoiation with genlteMACS dissociator</td></tr><tr><td>Halt protease and phosphatase inhibitor cocktial</td><td>ThermoFisher</td><td>78442</td><td>Component in the homogenization buffer</td></tr><tr><td>Laser doppler monitor</td><td>Moor</td><td>MOORVMS-LDF</td><td>Blood flow monitoring during tMCAO</td></tr><tr><td>LEGENDplex proinflammatory chemokine panel</td><td>Biolegend</td><td>740451</td><td>Multiplex bead array</td></tr><tr><td>LIVE/DEAD fixable near-IR stain</td><td>ThermoFisher</td><td>L34976</td><td>Use for dead cell exclusion during flow cytometric analysis</td></tr><tr><td>Ly6C-PE, clone HK1.4</td><td>Biolegend</td><td>128008</td><td>1:800 dilution</td></tr><tr><td>Ly6G-BV510, clone 1A8</td><td>Biolegend</td><td>127633</td><td>1:200 dilution</td></tr><tr><td>MCAO suture L56 reusable 6-0 medium</td><td>Doccol</td><td>602356PK10Re</td><td>tMCAO</td></tr><tr><td>MHC II-BV510, clone M5/114.15.2</td><td>Biolegend</td><td>107636</td><td>1:800 dilution</td></tr><tr><td>NK1.1-Percp/Cy5.5, clone PK136</td><td>Biolegend</td><td>108728</td><td>1:200 dilution</td></tr><tr><td>Siglec F-PE, clone E50-2440</td><td>BD Biosciences</td><td>552126</td><td>1:200 dilution</td></tr><tr><td>Silk suture thread, size 6/0</td><td>Fine Science Tools</td><td>18020-60</td><td>tMCAO</td></tr><tr><td>SomnoSuite anesthesia system</td><td>Kent Scientific</td><td>SS-01</td><td>Mouse anaesthetization for tMCAO</td></tr><tr><td>TCRb-BV510, clone H57-897</td><td>Biolegend</td><td>109234</td><td>1:200 dilution</td></tr><tr><td>Zirconia/silica beads, 2.3 mm</td><td>Biospec</td><td>11079125z</td><td>Beads for tissue homogenization</td></tr></tbody></table>

characterization of immune cells and proinflammatory mediators in the pulmonary environment

Immune cell expansion, activation, and trafficking to the lungs, which are controlled by the expression of multiple cytokines and chemokines, may be altered by severe brain injury. This is evidenced by the fact that pneumonia is a major cause of mortality in patients who have suffered from ischemic stroke. The goal of this protocol is to describe the use of multicolor flow cytometric analysis to identify 13 types of immune cells in the lungs of mice, including alveolar macrophages, interstitial macrophages, CD103+ or CD11b+ dendritic cells (DCs), plasmacytoid DCs, eosinophils, monocytes/monocyte-derived cells, neutrophils, lymphoid-derived&#160; T and B cells, NK cells, and NKT cells, following ischemic stroke induction by transient middle cerebral artery occlusion. Moreover, we describe the preparation of lung homogenates using a bead homogenization method, to determine the expression levels of 13 different cytokines or chemokines simultaneously by multiplex bead arrays coupled with flow cytometric analysis. This protocol can also be used to investigate the pulmonary immune response in other disease settings, such as infectious lung disease or allergic disease.

We recently reported that ischemic stroke induction in mice alters the immune cell composition of the lungs11. Specifically, transient cerebral ischemia increased percentages of alveolar macrophages, neutrophils, and CD11b+ DCs, while diminishing percentages of CD4+ T cells, CD8+ T cells, B cells, NK cells, and eosinophils in the pulmonary compartment. Moreover, cellular alteration corresponded to significantly diminished levels of multiple chemokines in the lungs. Described here is a method for t...

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Characterization of Immune Cells and Proinflammatory Mediators in the Pulmonary Environment

This protocol describes the use of flow cytometry to identify the changes in immune cell composition, cytokine profile, and chemokine profile in the pulmonary environment following transient middle cerebral artery occlusion, a murine model of ischemic stroke.

Immune cell expansion, activation, and trafficking to the lungs, which are controlled by the expression of multiple cytokines and chemokines, may be ...

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Immunology and Infection

3.2K Views.  West Virginia University. This protocol describes the use of flow cytometry to identify the changes in immune cell composition, cytokine profile, and chemokine profile in the pulmonary environment following transient middle cerebral artery occlusion, a murine model of ischemic stroke.

Video: Characterization of Immune Cells and Proinflammatory Mediators in the Pulmonary Environment

Analysis of Pulmonary Dendritic Cell Maturation and Migration during Allergic Airway Inflammation

Dendritic cells (DCs) are the key players involved in initiation of adaptive immune response by activating antigen-specific T cells. DCs are present in peripheral tissues in steady state; however in response to antigen stimulation, DCs take up the antigen and rapidly migrate to the draining lymph nodes where they initiate T cell response against the antigen1,2. Additionally, DCs also play a key role in initiating autoimmune as well as allergic immune response3.
DCs play an essential role in both initiation of immune response and induction of tolerance in the setting of lung environment4. Lung environment is largely tolerogenic, owing to the exposure to vast array of environmental antigens5. However, in some individuals there is a break in tolerance, which leads to induction of allergy and asthma. In this study, we describe a strategy, which can be used to monitor airway DC maturation and migration in response to the antigen used for sensitization. The measurement of airway DC maturation and migration allows for assessment of the kinetics of immune response during airway allergic inflammation and also assists in understanding the magnitude of the subsequent immune response along with the underlying mechanisms.
Our strategy is based on the use of ovalbumin as a sensitizing agent. Ovalbumin-induced allergic asthma is a widely used model to reproduce the airway eosinophilia, pulmonary inflammation and elevated IgE levels found during asthma6,7. After sensitization, mice are challenged by intranasal delivery of FITC labeled ovalbumin, which allows for specific labeling of airway DCs which uptake ovalbumin. Next, using several DC specific markers, we can assess the maturation of these DCs and can also assess their migration to the draining lymph nodes by employing flow cytometry.

We describe a strategy to monitor maturation and migration of pulmonary dendritic cells in response to ovalbumin in the setting of ovalbumin induced allergic airway inflammation. This strategy can be modified to assess migration of pulmonary dendritic cells in settings of infection.

Dendritic cells (DCs) are the key players involved in initiation of adaptive immune response by activating antigen-specific T cells. DCs are present in ...

A Method for Generating Pulmonary Neutrophilia Using Aerosolized Lipopolysaccharide

Acute lung injury (ALI) is a severe disease characterized by alveolar neutrophilia, with limited treatment options and high mortality. Experimental models of ALI are key in enhancing our understanding of disease pathogenesis. Lipopolysaccharide (LPS) derived from gram positive bacteria induces neutrophilic inflammation in the airways and lung parenchyma of mice. Efficient pulmonary delivery of compounds such as LPS is, however, difficult to achieve. In the approach described here, pulmonary delivery in mice is achieved by challenge to aerosolized Pseudomonas aeruginosa LPS. Dissolved LPS was aerosolized by a nebulizer connected to compressed air. Mice were exposed to a continuous flow of LPS aerosol in a Plexiglas box for 10 min, followed by 2 min conditioning after the aerosol was discontinued. Tracheal intubation and subsequent bronchoalveolar lavage, followed by formalin perfusion was next performed, which allows for characterization of the sterile pulmonary inflammation. Aerosolized LPS generates a pulmonary inflammation characterized by alveolar neutrophilia, detected in bronchoalveolar lavage and by histological assessment. This technique can be set up at a small cost with few appliances, and requires minimal training and expertise. The exposure system can thus be routinely performed at any laboratory, with the potential to enhance our understanding of lung pathology.

We describe a method for inducing neutrophilic pulmonary inflammation by challenge to aerosolized lipopolysaccharide by nebulization, to model acute lung injury. In addition, basic surgical techniques for lung isolation, tracheal intubation and bronchoalveolar lavage are also described.

Acute lung injury (ALI) is a severe disease characterized by alveolar neutrophilia, with limited treatment options and high mortality. Experimental models ...

Bronchoalveolar Lavage of Murine Lungs to Analyze Inflammatory Cell Infiltration

Bronchoalveolar Lavage (BAL) is an experimental procedure that is used to examine the cellular and acellular content of the lung lumen ex vivo to gain insight into an ongoing disease state.
Here, a simple and efficient method is described to perform BAL on murine lungs without the need of special tools or equipment. BAL fluid is isolated by inserting a catheter in the trachea of terminally anesthetized mice, through which a saline solution is instilled into the bronchioles. The instilled fluid is gently retracted to maximize BAL fluid retrieval and to minimize shearing forces. This technique allows the viability, function, and structure of cells within the airways and BAL fluid to be preserved.
Numerous techniques may be applied to gain further understanding of the disease state of the lung. Here, a commonly used technique for the identification and enumeration of different types of immune cells is described, where&#160;flow cytometry is combined with a select panel of fluorescently labeled cell surface-specific markers. The BAL procedure presented here can also be used to analyze infectious agents, fluid constituents, or inhaled particles within murine lungs.

The health status of the lung is reflected by the type and number of immune cells that are present in the bronchioles of the lung. We describe a bronchoalveolar lavage technique that allows the isolation and study of nonadherent cells and soluble factors from the lower respiratory tract of mice.

Bronchoalveolar Lavage (BAL) is an experimental procedure that is used to examine the cellular and acellular content of the lung lumen ex vivo to gain ...

An IL-8 Transiently Transgenized Mouse Model for the In Vivo Long-term Monitoring of Inflammatory Responses

Airway inflammation is often associated with bacterial infections and represents a major determinant of lung disease. The in vivo determination of the pro-inflammatory capabilities of various factors is challenging and requires terminal procedures, such as bronchoalveolar lavage and the removal of lungs for in situ analysis, precluding longitudinal visualization in the same mouse. Here, lung inflammation is induced through the intratracheal instillation of Pseudomonas aeruginosa culture supernatant (SN) in transiently transgenized mice expressing the luciferase reporter gene under the control of a heterologous IL-8 bovine promoter. Luciferase expression in the lung is monitored by in vivo bioluminescent image (BLI) analysis over a 2.5- to 48-h timeframe following the instillation. The procedure can be repeated multiple times within 2 - 3 months, thus permitting the evaluation of the inflammatory response in the same mice without the need to terminate the animals. This approach permits the monitoring of pro- and anti-inflammatory factors acting in the lung in real time and appears suitable for functional and pharmacological studies.

An IL-8 Transiently Transgenized Mouse Model for the In Vivo Long-term Monitoring of Inflammatory Responses

The method described here allows for the visualization of IL-8 promoter-dependent inflammation activation in the lungs of mice through non-invasive bioluminescence imaging (BLI). The same animal can be subjected to BLI multiple times for up to two months from the time of delivery of the luciferase reporter construct.

Airway inflammation is often associated with bacterial infections and represents a major determinant of lung disease. The in vivo determination of the ...

Noninvasive Sampling of Mucosal Lining Fluid for the Quantification of In Vivo Upper Airway Immune-mediator Levels

This protocol describes noninvasive sampling of undisturbed upper airway mucosal lining fluid. It also details the extraction procedure used prior to the analysis of immune mediators in fluid eluates for the study of the airway topical immune signature, without the need for stimulation procedures (often used by other techniques). The mucosal lining fluid is sampled on a strip of filter paper placed at the anterior part of the inferior turbinate and left for 2 min of absorption. Analytes are eluted from the filter papers, and the extracted protein-based eluates are analyzed by an electrochemiluminescence-based immunoassay, allowing for the high-sensitivity quantification of low- and high-level analytes in the same sample. We measured the in vivo levels of 20 preselected immune mediators related to specific immune signaling pathways in the upper airway mucosa, but the technique is not limited to that specific panel or sampling site. The technique was first implemented in 7-year-old children from the Copenhagen Prospective Studies on Asthma in Childhood2000 (COPSAC2000) cohort with allergic rhinitis. It was thereafter used in the longitudinal COPSAC2010 birth cohort, sampled at 1 month, 2 years, and 6 years of age and at instances of acute respiratory symptoms. We successfully obtained and analyzed samples from 620 (89%) of 700 1-month-old children; a few samples were below the assay detection limit (reported as the median (Inter-Quartile Range (IQR)). The number of samples below the detection limit (i.e.&#160;from 0 to the set point for the lower limit of detection) for each mediator was 29 (7.25 - 119.5). This technique enables the quantification of the in vivo airway mucosal immune profile from birth, can be applied longitudinally, and can be applied to studies on the effect of genetics and early-life environmental exposures, pathophysiology, endotyping, and monitoring of respiratory diseases, and development and evaluation of novel therapeutics.

Noninvasive Sampling of Mucosal Lining Fluid for the Quantification of In Vivo Upper Airway Immune-mediator Levels

This protocol describes a noninvasive technique for the sampling of undisturbed mucosal lining fluid from the upper airways. It can be used to perform the quantification of in vivo levels of protein mediators, such as cytokines and chemokines, in subjects of all ages.

This protocol describes noninvasive sampling of undisturbed upper airway mucosal lining fluid. It also details the extraction procedure used prior to the ...

Isolation and In Vitro Culture of Murine and Human Alveolar Macrophages

Alveolar macrophages are terminally differentiated, lung-resident macrophages of prenatal origin. Alveolar macrophages are unique in their long life and their important role in lung development and function, as well as their lung-localized responses to infection and inflammation. To date, no unified method for identification, isolation, and handling of alveolar macrophages from humans and mice exists. Such a method is needed for studies on these important innate immune cells in various experimental settings. The method described here, which can be easily adopted by any laboratory, is a simplified approach to harvesting alveolar macrophages from bronchoalveolar lavage fluid or from lung tissue and maintaining them in vitro. Because alveolar macrophages primarily occur as adherent cells in the alveoli, the focus of this method is on dislodging them prior to harvest and identification. The lung is a highly vascularized organ, and various cell types of myeloid and lymphoid origin inhabit, interact, and are influenced by the lung microenvironment. By using the set of surface markers described here, researchers can easily and unambiguously distinguish alveolar macrophages from other leukocytes, and purify them for downstream applications. The culture method developed herein supports both human and mouse alveolar macrophages for in vitro growth, and is compatible with cellular and molecular studies.

Isolation and In Vitro Culture of Murine and Human Alveolar Macrophages

This communication describes methodologies for isolation and culture of alveolar macrophages from humans and murine models for experimental purposes.

Alveolar macrophages are terminally differentiated, lung-resident macrophages of prenatal origin. Alveolar macrophages are unique in their long life and ...

Identification and Characterization of Immunogenic RNA Species in HDM Allergens that Modulate Eosinophilic Lung Inflammation

Environmental allergens such as house dust mites (HDM) are often in complex forms containing both allergic proteins that drive aberrant type 2 responses and microbial substances that induce innate immune responses. These allergen-associated microbial components play an important role in regulating the development of type 2 inflammatory conditions such as allergic asthma. However, the underlying mechanisms remain largely undefined. The protocol presented here determines the structural characteristics and in vivo activity of allergen-associated immunostimulatory RNA. Specifically, common allergens are examined for the presence of double-stranded RNA (dsRNA) species that can stimulate IFN responses in lungs and restrain&#160;the development of severe lung eosinophilia in a mouse model of HDM-induced allergic asthma. Here, we have included the following three assays: Dot blot to show the dsRNA structures in total RNA isolated from allergens including HDM species, RT-qPCR to measure the activities of HDM RNA in interferon stimulating genes (ISGs) expression in mouse lungs and FACS analysis to determine the effects of HDM RNA on the number of eosinophils in BAL and lung, respectively.

Environmental allergens such as house dust mites (HDM) often contain microbial substances that activate innate immune responses to regulate allergic inflammation. The protocol presented here demonstrates the identification of dsRNA species in HDM allergens and characterization of their immunogenic activities in modulating eosinophilic lung inflammation.

Environmental allergens such as house dust mites (HDM) are often in complex forms containing both allergic proteins that drive aberrant type 2 responses ...

Visualizing Lung Cellular Adaptations during Combined Ozone and LPS Induced Murine Acute Lung Injury

Lungs are continually faced with direct and indirect insults in the form of sterile (particles or reactive toxins) and infectious (bacterial, viral or fungal) inflammatory conditions. An overwhelming host response may result in compromised respiration and acute lung injury, which is characterized by lung neutrophil recruitment as a result of the patho-logical host immune, coagulative and tissue remodeling response. Sensitive microscopic methods to visualize and quantify murine lung cellular adaptations, in response to low-dose (0.05 ppm) ozone, a potent environmental pollutant in combination with bacterial lipopolysaccharide, a TLR4 agonist, are crucial in order to understand the host inflammatory and repair mechanisms. We describe a comprehensive fluorescent microscopic analysis of various lung and systemic body compartments, namely the broncho-alveolar lavage fluid, lung vascular perfusate, left lung cryosections, and sternal bone marrow perfusate. We show damage of alveolar macrophages, neutrophils, lung parenchymal tissue, as well as bone marrow cells in correlation with a delayed (up to 36-72 h) immune response that is marked by discrete chemokine gradients in the analyzed compartments. In addition, we present lung extracellular matrix and cellular cytoskeletal interactions (actin, tubulin), mitochondrial and reactive oxygen species, anti-coagulative plasminogen, its anti-angiogenic peptide fragment angiostatin, the mitochondrial ATP synthase complex V subunits, &#945; and &#946;. These surrogate markers, when supplemented with adequate in vitro cell-based assays and in vivo animal imaging techniques such as intravital microscopy, can provide vital information towards understanding the lung response to novel immunomodulatory agents.

Combined ozone and bacterial endotoxin exposed mice show wide-spread cell death, including that of neutrophils. We observed cellular adaptations such as disruption of cytoskeletal lamellipodia, increased cellular expression of complex V ATP synthase subunit &#946; and angiostatin in broncho-alveolar lavage, suppression of the lung immune response and delayed neutrophil recruitment.

Lungs are continually faced with direct and indirect insults in the form of sterile (particles or reactive toxins) and infectious (bacterial, viral or ...

Advanced Imaging of Lung Homing Human Lymphocytes in an Experimental In Vivo Model of Allergic Inflammation Based on Light-sheet Microscopy

Overwhelming tissue accumulation of highly activated immune cells represents a hallmark of various chronic inflammatory diseases and emerged as an attractive therapeutic target in the clinical management of affected patients. In order to further optimize strategies aiming at therapeutic regulation of pathologically imbalanced tissue infiltration of pro-inflammatory immune cells, it will be of particular importance to achieve improved insights into disease- and organ-specific homing properties of peripheral lymphocytes. The here described experimental protocol allows to monitor lung accumulation of fluorescently labeled and adoptively transferred human lymphocytes in the context of papain-induced pulmonary inflammation. In contrast to standard in vitro assays frequently used for the analysis of immune cell migration and chemotaxis, the now introduced in vivo setting takes into account lung-specific aspects of tissue organization and the influence of the complex inflammatory scenario taking place in the living murine organism. Moreover, three-dimensional cross-sectional light-sheet fluorescence microscopic imaging does not only provide quantitative data on infiltrating immune cells, but also depicts the pattern of immune cell localization within the inflamed lung. Overall, we are able to introduce an innovative technique of high value for immunological research in the field of chronic inflammatory lung diseases, which can be easily applied by following the provided step-by-step protocol.

The here introduced protocol allows characterization of the lung homing capacity of primary human lymphocytes under in vivo inflammatory conditions. Pulmonary infiltration of adoptively transferred human immune cells in a mouse model of allergic inflammation can be imaged and quantified by light-sheet fluorescence microscopy of chemically cleared lung tissue.

Overwhelming tissue accumulation of highly activated immune cells represents a hallmark of various chronic inflammatory diseases and emerged as an ...

Medicine

Flow Cytometric Analysis for Identification of the Innate and Adaptive Immune Cells of Murine Lung

The respiratory tract is in direct contact with the outside environment and requires a precisely regulated immune system to provide protection while suppressing unwanted reactions to environmental antigens. Lungs host several populations of innate and adaptive immune cells that provide immune surveillance but also mediate protective immune responses. These cells, which keep the healthy pulmonary immune system in balance, also participate in several pathological conditions such as asthma, infections, autoimmune diseases, and cancer. Selective expression of surface and intracellular proteins provides unique immunophenotypic properties to the immune cells of the lung. Consequently, flow cytometry has an instrumental role in the identification of such cell populations during steady-state and pathological conditions. This paper presents a protocol that describes a consistent and reproducible method to identify the immune cells that reside in the lungs of healthy mice under steady-state conditions. However, this protocol can also be used to identify changes in these cell populations in various disease models to help identify disease-specific changes in the lung immune landscape.

In this study, we present an effective and reproducible protocol to isolate the immune populations of the murine respiratory system. We also provide a method for the identification of all innate and adaptive immune cells that reside in the lungs of healthy mice, using a 9-color-based flow cytometry panel.

The respiratory tract is in direct contact with the outside environment and requires a precisely regulated immune system to provide protection while ...

Characterization of Immune Cells and Proinflammatory Mediators in the Pulmonary Environment

Summary

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Chapters in this video

Analysis of Pulmonary Dendritic Cell Maturation and Migration during Allergic Airway Inflammation

A Method for Generating Pulmonary Neutrophilia Using Aerosolized Lipopolysaccharide

Bronchoalveolar Lavage of Murine Lungs to Analyze Inflammatory Cell Infiltration

An IL-8 Transiently Transgenized Mouse Model for the In Vivo Long-term Monitoring of Inflammatory Responses

Noninvasive Sampling of Mucosal Lining Fluid for the Quantification of In Vivo Upper Airway Immune-mediator Levels

Isolation and In Vitro Culture of Murine and Human Alveolar Macrophages

Identification and Characterization of Immunogenic RNA Species in HDM Allergens that Modulate Eosinophilic Lung Inflammation

Visualizing Lung Cellular Adaptations during Combined Ozone and LPS Induced Murine Acute Lung Injury

Advanced Imaging of Lung Homing Human Lymphocytes in an Experimental In Vivo Model of Allergic Inflammation Based on Light-sheet Microscopy

Flow Cytometric Analysis for Identification of the Innate and Adaptive Immune Cells of Murine Lung