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

Summary

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 β and angiostatin in broncho-alveolar lavage, suppression of the lung immune response and delayed neutrophil recruitment.

Abstract

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, α and β. 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.

Introduction

Acute lung injury (ALI) is a crucial pathologic response of lungs to infectious or other harmful stimuli which is marked by simultaneous activation of coagulative, fibrinolytic and innate immune systems¹. Neutrophils promptly sense microbial as well as intracellular damage patterns through the Toll-like receptor (TLR) family^2,3,4. Neutrophils release preformed cytokines and cytotoxic granule contents, which can then cause collateral tissue damage. The ensuing alveolar damage is marred with secondary cell death leading to release of molecules such as adenosine triphosphate (ATP)⁵, thus setting in a vicious cycle of immune-dysregulation.

An unsolved problem in the understanding of ALI relates to the question of how the injury is initiated within the alveolar membrane. The electron transport complex V, F1F0 ATP synthase, is a mitochondrial protein known to be expressed ubiquitously, on cell (including endothelial, leukocyte, epithelial) plasma membrane during inflammation. The cell cytoskeleton which is comprised of actin and tubulin, harbors many cell shape and function modulating as well as mitochondrial proteins, respectively. We have recently shown that blockade of the ATP synthase by an endogenous molecule, angiostatin, silences neutrophil recruitment, activation and lipopolysaccharide (LPS) induced lung inflammation⁶. Thus, both biochemical (ATP synthase) and immune (TLR4) mechanisms might regulate the alveolar barrier during lung inflammation.

Exposure to ozone (O₃), an environmental pollutant, impairs lung function, increases susceptibility to pulmonary infections, and short low-levels of O₃ exposures increase the risk of mortality in those with underlying cardiorespiratory conditions^{7,8,9,10,11,12,13,14}. Thus, exposure to physiologically relevant concentrations of O₃ provides a meaningful model of ALI to study fundamental mechanisms of inflammation^7,8. Our lab has recently established a murine model of low-dose O₃ induced ALI¹⁵. After performing a dose and time-response to low O₃ concentrations, we observed that exposure to 0.05 ppm O₃ for 2 h, induces acute lung injury that is marked by lung ATP synthase complex V subunit β (ATPβ) and angiostatin expression, similar to the LPS model. Intravital lung imaging revealed disorganization of alveolar actin microfilaments indicating lung damage, and ablation of alveolar septal reactive oxygen species (ROS) levels (indicating abrogation of baseline cell signaling) and mitochondrial membrane potential (indicating acute cell death) after 2 h exposure to 0.05 ppm O₃¹⁵ which correlated with a heterogeneous lung ¹⁸FDG retention¹⁶, neutrophil recruitment and cytokine release, most notably IL-16 and SDF-1α. The take-home message from our recent studies is that O₃ produces exponentially high toxicity when exposed at concentrations below the allowed limits of 0.063 ppm over 8 h (per day) for human exposure. Importantly, no clear understanding exists on whether these sub-clinical O₃ exposures can modulate TLR4-mediated mechanisms such as by bacterial endotoxin¹⁷. Thus, we studied a dual-hit O₃ and LPS exposure model and observed the immune and non-immune cellular adaptations.

We describe a comprehensive fluorescent microscopic analysis of various lung and systemic body compartments, namely the broncho-alveolar lavage fluid (i.e., BAL) which samples the alveolar spaces, the lung vascular perfusate (i.e., LVP) that samples the pulmonary vasculature and the alveolar septal interstitium in the event of a compromised endothelial barrier, left lung cryosections, to look into resident parenchymal and adherent leukocytes left in the lavaged lung tissue, peripheral blood which represents the circulating leukocytes and the sternal and femur bone marrow perfusates that sample the proximal and distal sites of hematopoietic cell mobilization during inflammation, respectively.

Protocol

The study design was approved by the University of Saskatchewan's Animal Research Ethics Board and adhered to the Canadian Council on Animal Care guidelines for humane animal use. Six-eight week old male C57BL/6J mice were procured. NOTE: Euthanize any animals which develop severe lethargy, respiratory distress or other signs of severe distress before scheduled end point.

NOTE: Prepare the following: 27-18 G needle-blunted (will depend on the mouse tracheal diameter), appropriately sized PE tubing to fit the blunt needle (make a PE cannula for every mouse), cannula, 2 sharp scissors, 2 blunt forceps (small), 1 sharp forceps (small), 3 1 mL syringes, labelled microfuge tubes (for BAL, blood, vascular and bone marrow perfusate collection) and labelled vials (for tissue fixation), sample bags/cryovials for tissue collection, butcher's cotton thread roll (cut into adequate sized ligatures). All chemicals utilized for the experiments are indicated in the Table of Materials.

1. Ozone and LPS exposures for induction of murine lung injury

1. Ozone exposure
   1. Prepare a custom O₃ induction box. Add mouse feed, water bottle, enrichment toys and clean bedding in order and mimic the mice' housing environment.
      NOTE: These steps will ensure that the mice have free access to food and water when housed in the custom induction box and will help alleviate undue stress.
   2. Switch the O₃ calibrator/generator "ON" (placed towards the inlet port) to stabilize the UV lamp before hand (around 15 min before exposures) and connect with in-line O₃ monitor.
      NOTE: The O₃ monitor should read an average of 0.05 ppm, as sampled from the outlet at constant chamber air temperature (72 ±3 °F) and relative humidity (50±15%).
   3. For O₃ exposures, gently place mice in the custom induction box and continuously expose the mice to 0.05 ppm O₃ for 2 h.
2. Anesthesia and intranasal LPS administration
   1. Prepare 10 mg/mL stocks for ketamine and xylazine, separately.
   2. Prepare a cocktail by mixing 20 parts of ketamine and 1 part of xylazine. If the mouse weighs X g, inject (X/200) mL of ketamine/xylazine cocktail into the peritoneal cavity. For one mouse, prepare 1 mL of cocktail comprising of 1000 µL of the 10 mg/mL ketamine stock and 50 µL of the xylazine stock. Mix well by pipetting up and down in a microfuge tube.
      NOTE: The ketamine and xylazine stocks can be prepared a week in advance.
   3. Anesthetize the mice under light intraperitoneal (IP) ketamine (50 mg/kg)/xylazine (1 mg/kg) mix (e.g., for a 25 g mouse, inject 0.125 mL of the cocktail).
   4. Prepare a 1 mg/mL solution of LPS, in saline, and fill 50 µL of the solution in a pipette.
   5. Hold the mouse upright with its back/dorsal side on the palm, holding the ears with the same hand. Now, place 50 µL of the LPS solution¹⁸ gently outside nostrils (i.e., 25 µL on each nostril or 50 µL in one nostril; does not matter as long as the procedure is quick) and allow the mice to inhale the LPS solution.
   6. Instill control mice with 50 µL of sterile saline.
      Caution: Do not exceed more than 100 µL for instillation, which could be fatal. Too less of a volume (i.e., <50 µL) and there is risk of only nasal deposition.
   7. Following LPS administration, gently lay down the mice, either on their stomach or their back upon the mound of bedding with their head angled slightly downwards.
      NOTE: This orientation prolongs retention of the solution in the nasal cavity¹⁹.
   8. At 0, 2, 4, 24, 36 and 72 h after the exposure, anesthetize the mice i.p. with full dose of ketamine (200 mg/kg)/xylazine (4 mg/kg) mix (i.e., X/50 mL of the cocktail, where X is the mouse weight in grams or 0.50 mL for a 25 g mouse).
   9. Observe the mouse until it loses consciousness and the right reflex (i.e., does not turn back when laid in supine position). Now, check the mouse for depth of anesthesia, by monitoring the breathing (rhythmic chest excursions) and heart rate, which should fall noticeably, after 1-2 minutes.
   10. Next, check for pedal reflexes i.e., pinch either of the hind-limb digits and observe for retraction of the limb, as a reflex. If the mouse responds, top-up with additional 0.1 mL injections or more, as required.
       ​NOTE: To achieve deep anesthesia, bear in mind that certain strains or obese mice usually have a larger volume of distribution and thus can be easily over-dosed, if sufficient time if not allowed for full anesthesia (which can be around 10 minutes or more). Accordingly, some strains can be resistant to anesthesia and thus require more top-ups. This knowledge is acquired after many practical observations and experience with mice of different strains and age. As a few pilot experiments were also performed immediately after O₃ and LPS exposures (i.e., at the 2 h time-point), we also included some very early BAL cell analysis from those experiments to highlight the immediate effects of the combined O₃^{and LPS exposure.}

2. Sample collection

1. Place the mouse on surgical tray. Now, gently put lubricating eye ointment on both the eyes to avoid fluid loss and sanitize the mouse with 70% ethanol spray.
   1. Make small cuts through the upper and lower skin membranes with scissors, to expose the trachea and thorax.
   2. Make a cut just below the sternum and expose the heart.
2. Cardiac puncture
   1. Place a 25G needle attached to heparinized syringe into the right ventricle and draw blood by cardiac puncture.
      ​NOTE: Blood usually draws with minimal plunger draw, if the time from exposing the heart to cardiac puncture is under a minute. After collecting around 0.4-0.5 mL, one may need to pause for a few seconds before collecting the remaining blood. This is done to let the heart pump any remaining volume into the right ventricular chamber. By the end of the blood collection, heart stops to pump.
   2. Collect the blood and store in microfuge tubes for further processing.
3. Tracheostomy
   1. Carefully use tweezers/blunt forceps to clear off the tracheal region from overlying tissue. Use gloved hands more than the surgical instruments to avoid bleeding. If a lot of blood is oozing, there is risk of contaminating the BAL samples. Use cotton swabs, if necessary, although not preferred. Kimwipes are better alternatives.
   2. Now, cut through the ribcage to expose the lungs. Again, be cautious not to cut the sternum and the intercostal arteries or the ascending aorta; make smaller cuts while advancing through the rib cage)
   3. Pass a cotton ligature below the tracheal snip and leave as such for the time being.
   4. Snip the trachea at a suitable location in the distal 1/3^rd portion, pointing towards the lungs, to allow access for a 28 G tracheal cannula.
   5. Insert a 28 G PE cannula (a minimal length of 3-5 mm and distal end trimmed for ease of insertion) into the trachea. Watch out for the end of the trachea before bifurcation and then pull about a mm back to avoid sampling only single lobe.
   6. Firmly, tie cotton ligature to hold the cannula in place. Do not collapse the cannula.
4. Broncho-alveolar lavage (BAL)
   1. Fill 0.5 mL of PBS in a 1 mL syringe.
   2. Now gradually inject 0.5 mL of PBS into the cannula with the help of 1 mL syringe.
   3. After injecting PBS, aspirate out the syringe as long as it does not resist the suction.
      NOTE: If there is resistance while aspirating the BAL fluid out, that indicates collapse of the alveolar or bronchial tissue. In that case, pull back the cannula by a miniscule to detach the cannula form the walls of the tissue.
   4. Collect the aspirated fluid in a labelled microfuge tube and place on ice.
   5. Perform two more lavages in similar fashion collecting in the same vial (i.e., a total of 0.5 X 3 = 1.5 mL lavage).
   6. Measure the volume of collected BAL fluid. Lavage recovery should be close to 90%.
5. Lung vascular perfusate (LVP)
   1. Cut the descending thoracic aorta at a location between the thoracic and abdominal halves, to avoid any back-up of perfusate in the lungs while perfusing through the right ventricle.
   2. Blot the cavity near the cut aorta end, free of blood.
   3. Next, perfuse the lungs with 0.5 mL of room-temp heparinized saline injected through the right ventricle.
   4. Collect the vascular perfusate from the cavity at the cut end of the descending thoracic aorta, in a microfuge tube, placed on ice and measure its volume.
   5. Ligate the right bronchus proximal to its branching from trachea (with cotton thread).
6. In situ left lung fixation
   1. Connect a 1 mL syringe to tracheal cannula, which is long enough to insert through the previous tracheal incision.
   2. Draw back air in the syringe up to 0.6 mL mark.
   3. Then, fill the remaining syringe (up to the very end) with 2% paraformaldehyde at room temperature. Make sure that the plunger is ready to pop out without sucking back any air from the cannula.
   4. Now, affix the syringe with a scotch tape, to an upright container, measured up to 20 cm height.
   5. Gently, draw the plunger out to let the fixative flow towards the trachea. In case there are a few air bubbles in the cannula, gently place the plunger on top of the syringe and the fluid will start to flow after a few seconds.
   6. Let the left lung inflate to its total capacity in situ, for 5 minutes, from a 20 cm height forming a 20 cm water column.
   7. During this procedure, make sure to protect the right lung lobes from getting in contact with paraformaldehyde as this will affect the downstream assays.
   8. Place folded laboratory tissues to absorb any paraformaldehyde that might come in contact with the right lung lobes.
      ​CAUTION: Paraformaldehyde is highly toxic. Therefore, do not inhale or place contact to any exposed parts of the body. Exercise extreme caution while handling.
   9. During the time of paraformaldehyde instillation, sanitize the abdomen.
   10. Cut the right lung lobes off from the trachea making sure to remove any thread and immediately put the lobes in a labelled cryovial and drop it in liquid nitrogen for downstream molecular/biochemical cytokine analysis/RT-PCR/western blot analysis of lung homogenate.
   11. Carefully trim the left lung for any connective tissue or pleural membranes and dip it in 2% paraformaldehyde for 24 h at 4 °C.
   12. Embed the fixed lung in paraffin as per standard embedding protocol.
       Caution: Do not over-heat the lung samples.
   13. Cut the abdominal part and de-skin it from the pelvic (hip) bone.
   14. Separate out the left and right femur bones from the pelvic portion, in a saline filled petri dish kept on ice.
7. Sternal and femur bone marrow aspirate collection
   1. Clean the tissue and muscles off the bones by using laboratory tissues.
   2. Collect the ventral ribcage and sternum into a saline filled Petri dish kept on ice.
   3. Cut the distal and proximal tips of the sternal and femur bones.
   4. Perfuse the bones 4 times with 0.5 mL of saline, using well-fitted needles (24 to 28 G) on to 1 mL syringes, and collect the fractions from each bone into labelled tubes fitted with filters and placed on ice.
      NOTE: The needle gauge varies between animal size. After flushing, the femur bone should show up transparent.
   5. Once the samples have been collected, bag the mouse in a plastic bag and put in an animal carcass freezer for proper disposal, as per guidelines of the institutional animal facility.

3. Sample processing

1. Total (TLC) and differential (DLC) leukocyte counts
   1. Centrifuge peripheral blood, BAL, lung vascular perfusate and bone marrow (sternal and femur) samples for 10 min at 500 g.
   2. Collect the supernatants, flash freeze them and store them at -80 °C until further analysis.
   3. Reconstitute the cells in a minimum of 200 µL of PBS.
   4. Perform TLC by counting BAL, blood, lung vascular perfusate and bone marrow cells on a hemocytometer.
   5. Stain another 9 µL aliquot of BAL with a 1 µL mix of calcein green and red ethidium homodimer-1 to quantify live (green) cells due to intracellular esterase activity and any damaged BAL cells (in red) due to loss of plasma membrane integrity.
   6. Add 2% acetic acid to lyse RBCs, in a 1:10 ratio for blood TLC and 1:2 ratio for lung vascularperfusate TLC.
      CAUTION: Adjust the cell concentrations by diluting in PBS if the concentration is more than 1x10⁶ cells per mL (very important for the bone marrow samples).
   7. Centrifuge the cells to prepare cytospins on slides and stain as explained below for DLCs. Count a minimum of 100 cells for differential leukocyte cell counts (DLCs).
      NOTE: Typically, one can prepare two cytospins on one slide. And after 10-15 minutes of air drying, proceed with staining step.
   8. Split the collected BAL from three pilot mice per group into two cytospins each and stain for actin/tubulin and mitochondria with active oxidative phosphorylation (reduced mitotracker). Utilize the BAL from three more mice to split into two cytospins each, for NK1.1/Gr1/CX3CR1 and ATPβ/Ki-67/CD61/Angiostatin stained slides. Split the BAL from three more mice into two cytospins each to stain for ATPα/Ly6G and CX3CR1/Siglec-F.
2. Bronchoalveolar lavage (BAL) and Lung Vascular Perfusate (LVP) total protein quantification
   1. In order to quantify O₃ and LPS induced perturbations of the vascular barrier or the relative oncotic pressure in the two pulmonary compartments (i.e., the alveolar septal (interstitial) and lung vascular perfusate (vascular) compartments), measure total protein content in the collected fluids.
   2. Analyze the thawed supernatant fractions for their total protein concentration using a standard detergent resistant colorimetric assay.
   3. Bronchoalveolar lavage (BAL) and lung vascular perfusate (LVP) chemokine analysis
      1. Next, analyze the chemokines in BAL and lung vascular perfusate (LVP) supernatants using a 33-plex magnetic bead-based immunoassay. This will inform about the directionality of airway/interstitium vs vascular chemokine gradients established after combined exposures.
      2. Analyze the following panel of chemokines : CXCL13 (B-lymphocyte chemoattractant), CCL27 (IL-11 R-alpha-locus chemokine (ILC)), CXCL5 (epithelial-derived neutrophil-activating peptide 78 (ENA-78)), CCL-11 (eotaxin-1), eotaxin-2 (CCL-24), CX3CL1 (fractalkine), GM-CSF (CSF-2), CCL1, IFNγ (interferon gamma), IL-10 (interleukin-10), IL-16 (interleukin-16), IL-1β (interleukin-1 beta), IL-2 (interleukin-2), IL-4 (interleukin-4), IL-6 (interleukin-6), CXCL-10 (interferon gamma-induced protein 10 (IP-10)), CXCL11 (Interferon-gamma-inducible protein 9 (IP-9)), KC (keratinocyte chemoattractant), MCP-1 (monocyte chemoattractant protein-1), MCP-3 (monocyte chemoattractant protein-3), MCP-5 (monocyte chemoattractant protein-5), MDC (macrophage-derived chemokine (CCL22)), MIP-1α (macrophage inflammatory protein-1 alpha), MIP-1β (macrophage inflammatory protein-1 beta), MIP-2 (macrophage inflammatory protein-2), MIP-3α (macrophage inflammatory protein-3 alpha), MIP-3β (macrophage inflammatory protein-3 beta), RANTES (regulated on activation, normal T cell expressed and secreted (CCL5)), CXCL-16, CXCL-12/SDF-1alpha (stromal cell-derived factor 1), TARC (thymus and activation regulated chemokine (TARC)), TECK (Thymus-Expressed Chemokine (CCL25)) and TNFα (tumor necrosis factor alpha).

4. Cytospin staining and lung histology

1. Cytospin biochemical staining
   1. Encircle the cytospins with a hydrophobic pen to contain the chemicals for incubation during the procedure.
   2. Rehydrate the cytospins in PBS for 5 minutes.
   3. Fix the cytospin samples in 2% paraformaldehyde for 10 minutes, wash three times with PBS for 5 minutes each.
   4. Permeabilize with ice cold 70% acetone for 7 minutes and again wash three times with PBS for 5 minutes each.
   5. Stain for actin and tubulin or reduced Mitotracker (incubate with a mix of 2 µg/50 µL of Alexa 488 conjugated phalloidin shown in green, and 2 µg/µL of Alexa 555 conjugated mouse anti-α tubulin or reduced Mitotracker shown in red, respectively) for 15 minutes.
      ​NOTE: Use mouse IgG1 isotype control antibody, in a separate cytospin, to validate the tubulin staining protocol. Perform same steps as explained from 4.1.1 to 4.1.8, but for isotype controls.
   6. Remove the stains by gently tipping the mix from the slide. Incubate the slides with DAPI (4′,6-diamidino-2-phenylindole) for 5-10 min to stain nuclei.
   7. Wash the cytospins 3 times with PBS for 5 min each, coverslip with anti-fade mounting media and store overnight in dark at room temperature before imaging.
   8. Acquire images under a wide-field upright microscope equipped with a scientific camera. Ensure consistent camera exposure times set for the different fluorescent channels when imaging slides.
2. Cytospin immune-fluorescent staining:
   1. Encircle the cytospins with a hydrophobic pen to contain the chemicals for incubation during the procedure. Rehydrate the cytospins in PBS for 5 minutes.
   2. Fix the cytospin samples by incubating in 2% paraformaldehyde for 10 minutes, wash three times with PBS for 5 min each.
   3. Permeabilize with 0.1% cold triton X-100 for 2 minutes, wash three times with PBS for 5 min each.
   4. Next, Fc block for 15 minutes by incubating the cytospin with 1:50 dilution of the Fc block antibody stock (to ensure that the primary mouse antibody does not cross-react non-specifically with the mouse Fc antibodies).
   5. Tip the slides to remove the Fc block and change to 1% BSA for and incubate the cytospin with a mix of 3 primary antibodies of interest (refer to Table 1 for the various combinations) for 30 minutes.
      NOTE: Make sure to run isotype control antibodies (incubate with a mix mouse IgG1 and rat IgG2b kappa primary antibodies by performing steps 4.2.1 to 4.2.9 and replacing the antibodies with isotype controls) in parallel with corresponding secondary antibodies as discussed in our recent study²⁰.
   6. Wash 3 times with PBS for 5 min each. Incubate the cytospins with a mix of appropriately designed secondary antibodies (please refer to Table 1 for the details).
   7. Remove the stains by gently tipping the mix from the slide, incubate with DAPI (4′,6-diamidino-2-phenylindole) for 5-10 min to stain nuclei. Wash 3 times with PBS for 5 min each.
   8. Coverslip with anti-fade mounting media and store overnight in dark at room temperature before imaging.
   9. Acquire images under a wide-field upright microscope equipped with a scientific camera. Ensure consistent camera exposure times set for all the fluorophore channels when imaging slides.
3. Hematoxylin and eosin (H&E) histology
   1. Perform modified H&E staining³ on lung cryo-sections for all groups.
4. Image analysis
   1. Process and analyze image data (.tiff) files in Fiji ImageJ open software (https://imagej.net/Fiji/Downloads).
   2. Check the required parameters (Area, Perimeter, Integrated Density, Shape descriptors, Feret's diameter, Circularity, Display label) to be recorded under the Analyze tab and Set Measurements.
   3. Using the ROI manager, manually outline around 50-200 cells in the merged image panel using the "freehand selections" tool. Save the regions of interest (ROI) with Ctrl+T command and copy the saved ROIs for each cell over to all the fluorophore channels.
   4. Next press Ctrl+M to measure the pre-set parametersfor all the fluorescent channels (e.g., 405 nm (DAPI or ATPβ in blue), 488 nm (actin or NK1.1 or Ki-67 in green), 568 nm (tubulin or Gr1 or CD61 in red) and 633 nm (CX3CR1 or angiostatin in magenta)).
   5. Save the Results as .csv file under an appropriate name that represents your analysis. Copy the Results from the .csv file over to a spreadsheet and divide the fluorescence intensity (FI) (which is the Raw Integrated Density column from the Results file) of the stained molecule by that of DAPI or CD61 FI, as per staining design. These ratios are termed as "DAPI or CD61 normalized FI ratios".
   6. Next, plot these normalized ratios, circularity, cell perimeter and Feret diameter, to evaluate any change in cell size after the combined O₃ and LPS exposure.
   7. Use appropriate statistical software to check the normality of the collected data and to test the null hypothesis (please refer to the statistical analysis section below).
5. Statistical Analysis
   1. Express results as mean ± SEM. A minimum of three mice were used per group.
   2. For chemokine data analysis, adjust one-way ANOVA p-values for false discovery rate according to the Benjamini and Hoshberg correction.
   3. Analyze image parameters, cellularity, Feret diameter, perimeter, DAPI or CD61 normalized fluorescence intensity ratios by Mann-Whitney U test (for comparing two groups) or Kruskal Wallis test (for comparing multiple groups) as these data were not normally distributed.
   4. For rest of the imaging experiments, analyze the results using a one-way analysis of variance followed by Sidak's multiple comparisons. p-values <0.05 were considered significant.

Results

Combined O₃ and LPS exposure leads to systemic inflammation and bone marrow mobilization at 72 h: Cell counts in different compartments revealed significant changes in peripheral blood and the femur bone marrow total cell counts upon combined O₃ and LPS exposures. Although combined O₃ and LPS exposures did not induce any changes in the total BAL (Figure 1A) or LVP (Figure 1B

Discussion

The methods presented in the current study highlight the usefulness of multiple compartment analysis to study multiple cellular events during lung inflammation. We have summarized the findings in Table 2. We and many labs have extensively studied the murine response to intranasal LPS instillation, which is marked by rapid recruitment of lung neutrophils, which peaks between 6-24 h following which, resolution kicks in. And recently, we have shown that sub-clinical O₃ (at 0.05 ppm for 2 h) alone...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
33-plex Bioplex chemokine panel	Biorad	12002231
63X oil (NA 1.4-0.6) Microscope objectives	Leica	HCX PL APO CS (11506188)
Alexa 350 conjugated goat anti-mouse IgG (H+L)	Invitrogen	A11045
Alexa 488 conjugated goat anti-mouse IgG (H+L)	Invitrogen	A11002
Alexa 488 conjugated phalloidin	Invitrogen	A12370
Alexa 555 conjugated mouse anti-α tubulin clone DM1A	Millipore	05-829X-555
Alexa 568 conjugated goat anti-hamster IgG (H+L)	Invitrogen	A21112
Alexa 568 conjugated goat anti-rat IgG (H+L)	Invitrogen	A11077
Alexa 633 conjugated goat anti-rabbit IgG (H+L)	Invitrogen	A21070
Armenian hamster anti-CD61 (clone 2C9.G2) IgG1 kappa	BD Pharmingen	553343
C57BL/6 J Mice	Jackson Laboratories	64
Confocal laser scanning microscope	Leica	Leica TCS SP5
DAPI (4′,6-diamidino-2-phenylindole)	Invitrogen	D1306	aliquot in 2 µl stocks and store at -20°C
Inverted fluorescent wide field microscope	Olympus	Olympus IX83
Ketamine (Narketan)	Vetoquinol	100 mg/ml	Dilute 10 times to make a 10 mg/ml stock
Live (calcein)/Dead (Ethidium homodimer-1) cytotoxicity kit	Invitrogen	L3224
Mouse anti-ATP5A1 IgG2b (clone 7H10BD4F9)	Invitrogen	459240
Mouse anti-ATP5β IgG2b (clone 3D5AB1)	Invitrogen	A-21351
Mouse anti-NK1.1 IgG2a kappa (clone PK136)	Invitrogen	16-5941-82
Pierce 660 nm protein assay	Thermoscientific	22660
Rabbit anti-angiostatin (mouse aa 98-116) IgG	Abcam	ab2904
Rabbit anti-CX3CR1 IgG (RRID 467880)	Invitrogen	14-6093-81
Rat anti-Ki-67 (clone SolA15) IgG2a kappa	Invitrogen	14-5698-82
Rat anti-Ly6G IgG2a kappa (clone 1A8)	Invitrogen	16-9668-82
Rat anti-Ly6G/Ly6C (Gr1) IgG2b kappa (clone RB6-8C5)	Invitrogen	53-5931-82
Rat anti-mouse CD16/CD32 Fc block (clone 2.4G2)	BD Pharmingen	553142
Reduced mitotracker orange	Invitrogen	M7511
Xylazine (Rompun)	Bayer	20 mg/ml	Dilute 2 times to make a 10 mg/ml stock