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

All studies were conducted under the approval of the Institutional Care and Use Committee (IACUC) for Virginia Tech and in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals.

1. Intratracheal (i.t.) Inoculation of LPS Using Oropharyngeal Administration

1. Ensure that each animal is uniquely identified using either an ear punch, ear tag, or other institutionally approved method.
2. Record the baseline body weight and body temperature for each animal.
3. Prepare the working stock of LPS. Each mouse will receive a 50 µl dose of 1 mg/kg LPS in 1x phosphate buffered saline (PBS). Mock treated animals will receive a 50 µl dose of 1x PBS.
4. Prepare an appropriate chamber for the administration of isoflurane using the following drop method. Institutional guidelines vary regarding the use of isoflurane and other anesthetic agents and prior to initiating any studies, persons should contact their institution's Department of Animal Care/Welfare to insure the all guidelines are adequately met. If drop method isoflurane is not an option, other anesthetic agents are acceptable alternatives.
   1. In an appropriate safety cabinet, apply approximately 3 ml of isoflurane to a folded absorbent paper towel and place in the bottom of a 500 ml beaker.
   2. Place a small piece of aluminum foil on top of the paper towel and cover the beaker with a transparent lid.
5. Prepare the tools and reagents that will be utilized during the i.t. inoculation. Position the rubber band that will secure the animals head on the intubation stand. This procedure will require a pair of straight forceps, a pair of angled or curved forceps, and a p200 pipette with tips.
6. Load 50 µl of LPS into the pipette tip and place in a readily accessible location beside the intubation stand.
7. Anesthetize the mouse by placing the animal into the isoflurane chamber and covering the chamber with the transparent lid. Observe the animal's breathing patterns, which will be rapid and shallow when first placed in the chamber. The animal will be sufficiently anesthetized when breathing rates approach 1 breath/2 sec, which is usually reached within 30 sec of placing in the chamber. The mouse should be anesthetized as stated in the approved animal protocol using drop method isoflurane.
8. Remove the anesthetized mouse from the chamber and suspend the animal on the intubation stand by its front incisors. Insure that the animal is securely restrained and the mouth and tongue are accessible.
9. Gently secure the tongue with the straight forceps. Grasp the tongue with the angled forceps and gently pull it out of the mouth until slight resistance is felt. This action is sufficient to block the epiglottis and gain access to the trachea.
10. While continuing to hold the tongue in the extended position, administer the 50 µl dose of LPS into the back of the throat and immediately cover the animal's nostrils with a gloved finger. The LPS will be visible in the back of the throat until the mouse inhales.
11. Continue to cover the nostrils and hold the tongue extended for 5-10 additional breaths.
12. Remove the mouse from the intubation stand and place the animal back into a new cage until it recovers from the anesthesia.
13. Monitor surrogate markers associated with disease progression. Body weight and body temperature should be monitored every 4-8 hr for the duration of the study. Likewise, evaluate behavioral characteristics and clinical symptoms associated with increased morbidity.
14. To evaluate disease progression, harvest mice at specific time-points following the LPS exposure. Typical time-points for harvest following LPS exposure include 0, 6, 12, 18, 24, and 48 hr. To evaluate recovery and survival, evaluate the mice for 7 days post-inoculation.

2. Serum and Bronchoalveolar Lavage Fluid (BALF) Collection

1. Sacrifice the mouse using an institutionally approved method.
2. Place the animal on its back and secure it to a necropsy board, ideally 0.5 in (1.27 cm) in height.
3. Wet the entire mouse with 70% ethanol.
4. Using a 1 ml syringe with a 27 G needle, conduct a cardiac puncture prior to making any incisions. It should be possible to withdraw 500-800 µl of whole blood from a single mouse.
5. Remove the needle from the syringe and transfer the whole blood to a prelabeled 1.5 ml microcentrifuge tube. For serum collection, allow the whole blood to coagulate for at least 30 min at room temperature. In addition to serum collection, this procedure can also be modified by incorporating an anti-coagulant in the tube and syringe, prior to collecting the whole blood, to study circulating immune cells.
6. Make a horizontal incision across the length of the peritoneal cavity and a vertical incision from the peritoneal cavity to the lower jaw of the mouse.
7. Using forceps, grasp both sides of the incision and gently pull the skin away from the underlying peritoneal and thoracic cavities.
8. Make a large incision along the length of the peritoneal cavity from the genitals to the sternum, taking care to avoid cutting the diaphragm. Gently shift the intestines in the peritoneal cavity to allow access to one of the kidneys.
9. Cut the renal vein leading to the kidney to serve as a drainage point for perfusion.
10. Nick the diaphragm using scissors, taking care to avoid the lungs and heart, to expose the left side of the heart.
11. Manually perfuse the heart using a 10 ml syringe with a 27 G needle and 1x PBS solution. Avoid applying excessive force during the perfusion to ensure that the saline is not forced into the lungs. The remaining blood should drain from the portal vein incision.
12. Carefully cut the ribcage along the sternum using blunt/blunt scissors, taking care to avoid cutting the heart and lungs. Accidentally cutting the lungs during this procedure will significantly reduce the amount of BALF collected and will result in an inability to properly inflate the lungs with fixative.
13. Gently isolate the heart and lungs away from the ribcage using forceps. Carefully cut each section of the ribcage using blunt/blunt scissors as close to the spine as possible and fully remove both sections.
14. Separate the salivary glands using forceps or remove them using scissors, to expose the trachea.
15. Carefully cut the muscles that overlay the trachea using scissors. It is essential that the trachea not be cut during this process.
16. Using scissors, separate the collarbone overlying the trachea.
17. Gently grasp the thymus using forceps and lift the tissue away from the heart. Remove the thymus using scissors, while taking care to avoid cutting the heart or lungs.
18. Make a small horizontal incision in the trachea 1-2 rings below the larynx using angled scissors (45-90º angle, sharp/sharp). The incision should be large enough to firmly secure the cannula.
19. Insert the cannula so that the tapered end extends 2-3 tracheal rings below the incision and secure the cannula in place using a silk suture. Ensure that the suture passes between the trachea and the esophagus and is pulled tight on the cannula.
20. Fill a 1 ml syringe with 1 ml of Hanks Buffered Saline Solution (HBSS) and gently insert its luer into the cannula.
21. Using a slow, but constant motion, inject ~900 µl into the trachea, inflating the lungs, and immediately withdraw the HBSS. Place the recovered BALF in a 15 ml conical tube.
22. Repeat this lavage 2 additional times to collect approximately 3 ml of BALF for future analysis. Store the BALF on ice or at 4 ºC until ready for use.

3. Histopathology Preparation

1. Insert a 10 ml syringe into the lung inflation stand. Assemble the tubing to the luer, the luer to the stopcock, and the stopcock to the syringe. Ensure the stopcock is in the closed position. The 10 ml syringe will serve as a gravity flow reservoir. The reservoir should be attached to the inflation stand 4.75 in above the animal and the top of the reservoir should extend 10.5 in above the animal.
2. Fill the syringe with neutral buffered formalin solution. Ensure that the syringe is completely filled with the fixative.
3. Place an absorbent towel under the open end of the tubing and open the stopcock to allow the fixative to fill the tubing. Once the fixative begins flowing from the tubing, immediately close the stopcock and refill the syringe to the top with fixative. It is important that the syringe be completely filled to provide the appropriate amount of pressure for lung inflation.
4. Gently pass a second piece of suture thread under the trachea, approximately 1-2 tracheal rings below the end of the cannula.
5. Tie a single loose knot in the suture; however, do not pull the knot tight.
6. Insert the tubing from the mouse inflation stand into the cannula.
7. Open the stopcock and allow the lungs to fill by gravity inflation.
8. Once the lungs reach their maximum inflation level, pull the suture tight and tie a second knot.
9. Close the stopcock and remove the tubing from the cannula.
10. Gently remove the cannula from the trachea by grasping the suture with a pair of forceps and holding the cannula with fingers. Firmly pull the suture down towards the thoracic cavity and the cannula back towards the mouse's nose.
11. Gently grasp the trachea or the sutures with forceps and lift the trachea away from the neck cavity.
12. Sever the trachea caudal to the tied suture using blunt scissors.
13. Once the trachea is free from the underlying tissues, begin to pull the trachea away from the mouse. Gently lift the lungs out of the thoracic cavity.
14. Carefully cut the connective tissue holding the lungs in the thoracic cavity. Continue pulling the trachea away from the mouse's body and apply steady upward force while cutting the underlying connections. Take care to avoid cutting the lungs. Note that the heart should be left attached to the lungs using this method.
15. Once the lungs have been removed, place them in 10 ml of buffered formalin.
16. Remove a section of mouse tail for genotyping.
17. Dispose of the mouse carcass following the appropriate institutional guidelines.

4. Cytokine Evaluation

1. Centrifuge the whole blood, collected under step 2.5, at 12,000 x g for 5 min to isolate the serum. Transfer the serum to a prelabeled 1.5 ml microcentrifuge tube and store at -80 °C.
2. Evaluate the serum cytokine levels by ELISA or another similar assay. Dilute the serum 1:5 - 1:20 in an appropriate dilution buffer, depending on the assay, following the manufactures instructions. These dilutions should be empirically determined prior to running the bulk of the samples.
3. Due to the low volume of serum collected, reduce the sample volume loaded onto the ELISA plate by half. For example, most commercial ELISAs utilize 100 μl volumes of standards and samples. To conserve specimens, load 50 μl of standards and diluted serum per well.
4. Centrifuge the BALF in a refrigerated tabletop centrifuge at 200 x g for 5 min. Transfer the cell free supernatant into two prelabeled 1.5 ml microcentrifuge tubes and store at -80 °C.
5. Evaluate BALF cytokines by ELISA or another similar assay without diluting.
6. Store unused serum and BALF at -80 °C.

5. Differential Staining and BAL Cellularity Evaluation

1. Following BALF centrifugation and complete HBSS removal, lyse the pelleted red blood cells using hypotonic saline. Resuspend the cells in 900 μl of distilled water. Immediately add 100 μl of 10x PBS.
2. Individually lyse each sample. If samples contain excessive amounts of red blood cells, the cells can be pelleted in the table top centrifuge at 400 x g for 5 min and the red blood cell lysis protocol can be repeated.
3. Determine the total BALF cellularity in the 1 ml suspension using a hemacytometer under 10X or 20 X magnification with Trypan Blue staining. Evaluate and present these data as cells/ml.
4. Prepare materials for the cytospin and differential staining. Label standard microscope slides using a pencil or solvent resistant pen. Secure the slides into a cytospin bracket and funnel. Secure the slide assembly in the cytospin rotor.
5. Cytospin 150 μl of BALF at 100 x g for 5 min. If the cell density is too great to effectively evaluate cell morphology, reduce the volume spun down onto the slides. Allow the slides to air dry overnight.
6. Differential stain the slides following the manufactures protocols. Allow the slides to air dry overnight. Coverslip the slides using Permount and evaluate the slides using a microscope equipped with a 20X and 40X objective.
7. Harvest the remaining cells for subsequent analysis, such as FACS, electron microscopy, confocal microscopy, RNA extraction for gene expression evaluation, and/or protein extraction for western blot.

6. Histopathology Evaluation

1. Prepare the lungs for histopathology evaluation. After 24-48 hr of formalin fixation, ventrally orient the whole inflated lungs and embedded in paraffin. Cut the resultant blocks to expose the main conducting airway.
2. To improve scoring accuracy, position the lungs in the same position and trim each block to yield the maximum longitudinal visualization of the intrapulmonary main axial airway. From this point, cut 5 micron serial sections and stain with hematoxylin and eosin (H&E). Additional sections can be cut and prepared for in situ hybridization using standard protocols.
3. Evaluate histopathology using a semi-quantitative scoring system based on the following inflammatory parameters, which are scored between 0 (absent) and 3 (severe): mononuclear and polymorphonuclear cell infiltration; airway epithelial cell hyperplasia and injury; extravasation; perivascular and peribroncheolar cuffing; and the percent of the lung involved with inflammation. Lung histopathology should be evaluated by an experienced pathologist.
4. Average all of the parameter scores to generate a total histopathology score or use individual scores to quantify specific aspects of disease progression. Conduct all scoring in a double blind fashion, with reviewers blinded to both genotype and treatment. This scoring system has been previously described^6,7,10.