Adoptive Transfer of IL-33-Stimulated Macrophages into Bleomycin-Induced Mouse Models to Study Their Effect on Idiopathic Pulmonary Fibrosis In Vivo

All experiments were performed in accordance with the Guide for the Care and Use of Laboratory Animals. All the animal experiments were approved by the Experimental Animal Welfare Ethics committee of Jiangnan University (JN No. 20211¹³⁰m1720615[501]).

NOTE: In total, 10 male C57BL/6 mice aged 6-8 weeks old and weighing 20-25 g were used in this study. The three experimental groups in the study included three recipient mice each, and one host mouse was used for the IM isolation.

1. Depletion of mouse pulmonary macrophages

1. Take out the vial of clodronate liposomes (see Table of Materials) from the refrigerator 30 min in advance, warm it to room temperature, and invert it several times to ensure uniform mixing.
   NOTE: Clodronate liposomes encapsulate the hydrophilic dichloromethylene bisphosphonate molecules. When these liposomes are consumed by the macrophages, the clodronate is gradually released by the action of the lysosome phosphatase, and its accumulation consequently triggers cell apoptosis and the depletion of macrophages⁷.
2. Anesthetize mice with 3% isoflurane. Check the depth of anesthesia by pinching one foot to check consciousness. Apply veterinary ointment to both eyes to prevent dryness under anesthesia.
3. Aspirate 60 µL of clodronate liposomes using a pipette with a sterile suction tip. Administer the drug dropwise into the nasal cavity of the anesthetized mouse, such that when the mouse inhales, the drug is inhaled into the trachea. After each drop, make sure the mouse inhales the drug completely and is breathing evenly. Administer liposomes containing PBS alone as a control⁸ (Figure 2).
4. Place the mice on a 38 °C constant temperature table for rewarming to speed up their recovery. Monitor the mice until they wake up. After the mice recover well and achieve sternal recumbency, transfer them to the cage.
5. Use mice with depleted alveolar macrophages 2 days after the treatment with clodronate liposomes as the recipient mice for the transfer experiment.
   ​NOTE: In this study, the depletion of alveolar macrophages was confirmed by checking the expression of macrophage markers as described earlier^8,9 following the administration of clodronate liposomes by inhalation (see Supplementary Figure 1).

2. Isolation and culture of IMs

1. Anesthetize the host mouse with an intraperitoneal injection of Ketamine (120 mg/kg) and Xylazine (16 mg/kg). Confirm the depth of anesthesia via loss of the toe pinch reflex. Apply veterinary ointment to both eyes to prevent dryness under anesthesia. Then, disinfect the skin of the anesthetized mouse with 75% alcohol and iodine. Use scissors to cut through the skin and expose the cardiopulmonary tissue
2. Aspirate 10 mL of 1x PBS in a syringe with a 20 G needle and insert the tip of the needle into the right atrium of the mouse (Figure 3A). Cut the inferior vena cava of the mouse with surgical scissors, and then manually perfuse the mouse with PBS at a constant speed (10-20 mL/min) until the lung tissue turns white. Excise the lung tissue and transfer it into ice-cold PBS in a culture dish (Figure 3B).
3. Cut the lung tissue into fragments of approximately 2 mm x 2 mm x 2 mm. Then, add 15 mL of Dulbecco's modified Eagle's medium (DMEM) containing 1% collagenase A to the lung tissue to isolate the macrophages. Incubate at 100 rpm for 30 min on a 37 °C shaking table.
4. Aspirate the lung tissue suspension through a 10 mL syringe at least 20x to make it as fine as possible. Filter this suspension through a 40 µm cell strainer. Centrifuge the filtrate at 400 x g for 10 min and discard the supernatant.
5. Lyse the red blood cells in the pellet by adding 3 mL of RBC lysis buffer (see Table of Materials) on ice for 2-3 min.
6. After centrifugation at 150 x g for 5 min, resuspend the cell pellet with 10 mL of DMEM (containing 100 U/mL penicillin and 100 µg/mL streptomycin). Count the cells using a hemocytometer.
7. Seed the cells into 10 cm adherent cell culture dishes at 2 x 10⁷ cells/dish, and incubate for 1 h at 37 °C and 5% CO₂. This allows the lung IMs to adhere to the dish⁵. After 1 h, aspirate the supernatant and floating cells, and add 10 mL of fresh complete culture medium (DMEM containing 10% FBS, 100 U/mL penicillin, and 100 µg/mL streptomycin). Incubate for more than 8 h or overnight.
8. Determine the purity of the obtained IMs using flow cytometry with staining for F4/80 and CD11c markers, as described earlier⁵ (see Supplementary Figure 1).

3. Adoptive transfer of IMs into the lung alveoli

1. Replace the culture medium from the plate containing the isolated pulmonary IMs (step 2.7) with a complete culture medium containing 10 ng/mL IL-33 for stimulating the IMs. Incubate for 24 h at 37 °C and 5% CO₂. Use 1x PBS in place of IL-33 as a control.
2. Dissociate the pulmonary IMs by treating them with 1 mL of 0.25% trypsin for 5 min. Then, quench the digestion by adding 3 mL of fresh culture complete medium, and harvest the pulmonary macrophages by centrifuging the cell suspension at 150 x g and 4 °C for 5 min. Count the cells using a hemocytometer, and resuspend the cells in PBS to a final concentration of 5 x 10⁵ cells/50 µL.
3. Anesthetize the recipient mice with 3% isoflurane gas. Wait for the mouse to become unconscious (as in step 1.2), and then fix the limbs of the anesthetized mouse on a vertical plate with medical tape. Gently pull the mouse's tongue to one side using a cotton swab.
4. Turn on the intubation lamp and shine it on the throat of the mouse so that the trachea can be seen. Check that the trachea is close to the base of the tongue, the esophagus is near the back of the neck, but the opening of the esophagus is not visible during the operation.
5. Use the intubation lamp (22 G) to help intubate the mouse. Guide the indwelling needle cannula into the trachea with a guide wire (0.4 mm diameter). Remove the guide wire and push the cannula into the mouse trachea to complete the intubation.
6. After the cannula is observed to enter the trachea, inject 50 µL of the cell suspension (containing 5 x 10⁵ IMs) into the recipient mouse through the trachea.
7. Place the mice on a 38°C constant warming pad for rewarming to speed up the recovery. Monitor the mice until they wake up. After they recover well and achieve sternal recumbency, transfer them to the cage.
8. After 24 h, administer bleomycin (BLM) via the trachea (using the procedure in steps 3.3-3.5) at a dose of 1.4 U/kg body weight to induce IPF. Administer an equal volume of saline to the control group.
9. After 21 days, determine the degree of severity of pulmonary fibrosis for the different groups of mice that were adoptively transferred with PBS or the IL-33-stimulated IM suspension by evaluating the expression of markers for fibrosis and the Ashcroft scores¹⁰.
   1. Extract the total RNA from the lung tissue using an RNA extraction reagent (see Table of Materials).
      NOTE: Quantitative analysis was carried out with a microplate reader. If the OD260/OD280 ratio is 1.8-2.0, the RNA purity is high. All the samples were stored at −80 ° C.
   2. Perform fluorescence quantitative PCR to evaluate the expression of α-smooth muscle actin (SMA) and fibronectin using specific primers.
      ​NOTE: The primers used for α-SMA were forward 5'- GACGCTGAAGTATCCGATAGAACACG-3' and reverse 5'-CACCATCTCCAGAGTCCAGCACAAT-3', and the primers used for fibronectin were forward 5'-TCTGGGAAATGGAAAAGGGGAATGG-3' and reverse 5'-CACTGAAGCAGGTTTCCTCGGTTGT-3'.
10. Perform immunohistochemistry as described below.
    1. Dehydrate the left lung, embed it, and slice it with a paraffin slicer to a thickness of 4 µm.
    2. Place the tissue sections onto slides and bake the slides in an oven at 65-70 ° C for 30 min to 1 h. Dewax the slides with a decreasing percentage of alcohol (i.e., 100%, 95%, 90%, 80%, and 70% alcohol) for 5 min.
    3. Stain the slides in hematoxylin dye solution (analytically pure) for 5 min. Then, wash the slides with tap water. Soak the slides in 1% hydrochloric acid alcohol for 3 s, wash off the floating color, and soak in flowing water for 5 min. The sections will turn blue.
    4. Immerse in eosin solution for 10 s to 1 min (determine the dyeing time according to the color), wash with tap water, and place in 70%, 80%, 95%, 100%, and 100% alcohol, respectively, for 5 min each. Then place, the slides in xylene I and xylene II solutions for 3 min. After drying, observe and take photos under the vertical microscope with neutral adhesive sealing film.
    5. Perform Ashcroft scoring on the sections to indicate the severity of pulmonary fibrosis¹⁰. Perform statistical analysis of the data using a t-test of two independent samples.