Murine Double Hit Model for Neonatal Cardiopulmonary Diseases: Bronchopulmonary Dysplasia (BPD) and Pulmonary Hypertension Associated with BPD

Materials and Reagents

1. Ultra-fine 31G insulin syringes 0.5 mL (Becton Dickinson, catalog number: 32281)

2. Insyte^TM Autoguard^TM winged, 24G × 0.75” (Becton Dickinson, catalog number: 381212)

3. Microlance^TM hypodermic needle 30G × 0.5 (Becton Dickinson, catalog number: 304000)

4. 10 µL gastight syringe, cemented needle, 26s gauge, 2 in., point style 2 (Hamilton, catalog number: 1701)

5. C57BL/6J mice

6. Lipopolysaccharide (LPS) from Escherichia coli O127:B8; purified by phenol extraction (Sigma, catalog number: L3129)

7. Sodium chloride intravenous infusion BP (water for injection) (Pfizer, catalog number: AUST R 49280)

8. Oxygen gas

9. Isoflurane (AbbVie, Forthane^®, catalog number: AUST R 29656, store below 30 °C)

10. Paraformaldehyde (PFA) powder, 95% (Sigma-Aldrich, catalog number: 158127)

11. PBS (ThermoFisher Scientific, Gibco^TM, catalog number: 14190250)

12. Cotton tip applicators (BSN Medical, catalog number: 7505-0)

13. Gauze

14. Liquid nitrogen (N₂)

15. Wet ice

16. Polyester thread (sutures) (Birch, catalog number: 004601)

17. 1.7 mL microtubules (Axygen, catalogue number: MCT-175-C)

Equipment

1. Custom plexiglass gas chamber [approximate dimensions: 40 × 125 × 30 (L × W × H) cm]

2. Bear cub BP2001 infant ventilator (Bear Medical Systems, Type #240)

3. Portable oxygen analyzer (Teledyne, model: AX300-I)

4. Oxygen sensor (Maxtec, model: R109P09)

5. Power lab/16SP (ADinstruments, model: ML795)

6. Heated respiratory humidifiers (Fisher and Paykel, model: MR730)

7. Autofill humification chamber (Fisher and Paykel, model: MR290)

8. Temperature pod (ADinstruments, model: ML312)

9. Pod expander (ADinstruments, model: ML305)

10. Compact scale, 510 g × 0.1 g (A&D Weighing, model: HT-500)

11. Gravity perfusion apparatus

12. Forceps, tip width 0.5 mm, length 10 cm (Fine Science Tools, catalog number: 11150-10)

13. Scissors (Precision Medical Specialties, model: E25-500)

14. Foam dissection tray

15. Liquid nitrogen container (Nalgene^®, catalog number: 4150-2000)

16. -86 °C freezer 690 L (Froilabo, model: BMEVO69086G)

Software

1. Chart 5 Pro v5.5.1 (ADInstruments, www.ADInstruments.com)

Procedure

Murine double hit model of cardiopulmonary disease

1. Experimental conditions (Figure 2)

   
   

   
   Figure 2. Setup of an experimental chamber. A custom plexiglass chamber is connected to a humidifier and ventilator. Humidified oxygen gas with a fraction of inspired oxygen (FiO₂) of 0.65 is passed through the hyperoxia chamber at a flow rate of 10 L/min (air chamber has a FiO₂ of 0.21). Power lab recording system records temperature, humidity, and oxygen levels in chambers. Humidity and temperature are kept constant at 50%–60% and 22 °C, respectively. Light is cycled in a 12 h day/night rhythm. Chamber setup included below the schematic for comparison.

   
   

2. Time mating

   1. Use C57BL/6J mice.

   2. Breeders are housed in enhanced specific pathogen free 2 conditions.

   3. Preparation of female breeders:

      1. Select 6–14-week-old females for consideration.

         1. Younger females might not be sexually mature.

         2. Older females mate less reliably.

      2. One to two weeks prior to timed mating, house females together.

         Group housed females to synchronize their estrous cycles. This phenomenon is known as the “Lee-Boot effect” (Van Der Lee and Boot, 1955).

      3. Three days prior to timed mating, place material from a stud male cage into the female cages.

         The exposure of group housed female mice to urine from a male mouse increases the chance of ovulation on the third night following exposure. This phenomenon is known as the “Whitten effect” (Whitten, 1956).

   4. Preparation of male breeders:

      1. Select 6–14-week-old males.

         Preference proved stud males as sexual experience improves male reproductive performance (Swaney et al., 2012).

      2. One to two weeks prior to timed mating, separate the stud males.

         This allows the sperm count to increase and libido to recover from any previous mating.

   5. Timed mating:

      1. Place 1–2 females together with a stud male in a cage.

      2. Check for viscous vaginal plugs in the mornings.

         1. Components of the male ejaculate coagulate to form a hard plug that blocks the opening of the vagina and is an indicator of copulation. However, 37.1% of females that plug do not become pregnant (Heyne et al., 2015).

         2. A positive plug is considered embryonic age (E)0.5 if pregnancy occurs.

      3. After 48 h together, separate breeding pairs.

         
         

3. Early pregnancy detection

   1. Monitor body weight gain of females.

      Pregnant mice gain on average 3.49 g from E0 to E7–10 (Heyne et al., 2015).

   Or
   

   2. Palpate the pregnant dam’s abdomen with the thumb and forefinger at E12.

      A positive pregnancy feels like a string of beads.

      
      

4. Antenatal inflammatory hit 1 at E14

   1. Intraperitoneal injection of lipopolysaccharide (LPS) to the dam:

      1. Determine the dam’s body weight.

      2. Prepare LPS injection.

         1. Reconstitute LPS powder in water for injection.

         2. Final concentration injected is 150 µg/kg.

         3. Final volume injected is 1 µL/g.

      3. Inject LPS into the lower right quadrant of the dam at a 45° angle with an insulin syringe (Figure 3).

         Adverse events associated with i.p. injection are related to errors in the placement of the injection. Ensure that the needle passes through the skin, muscle, and peritoneum of the abdomen without penetrating any abdominal organs.

      4. Monitor dams closely to ensure welfare of animals.

         If miscarriage occurs, humanely euthanize the dam and perform a postmortem.

         
         

   
   Figure 3. Intraperitoneal injection site for LPS on day 14 of gestation in the pregnant dam

   
   

5. Postnatal inflammatory hit 2

   1. Dams deliver pups naturally at E19–21 in the saccular stage of lung development (Backstrom et al., 2011).

   2. The saccular stage is most susceptible to lung injury in mice (Backstrom et al., 2011), comparable to a premature human baby at 24–38 weeks gestation (Swaney et al., 2012).

   3. Within 24 h of birth, randomly assign pups and dams to either:

      1. Hyperoxia (FiO₂ of 0.65)

         Develop cardiopulmonary disease

      2. Normal room air (FiO₂ of 0.21)

         Control animals

   4. Set up foster pairs to avoid oxygen toxicity in the dam; pair one dam with pups in a hyperoxia cage with a counterpart dam and pups in an air cage.

      In case of an odd number of deliveries:

      1. Allocate the extra dam to an air group.

      Or

      2. Distribute the pups from the extra litter into the other litters under the experiment.

   5. Choose experimental endpoints to determine different readouts.

      1. Three days of 65% hyperoxia

         Pulmonary immunity:

         Postnatal exposure of pups to hyperoxia induces an immediate IL-1 and type 2 cytokine pulmonary immune response, which is most evident within the first three days of life. Accordingly, we see increased pulmonary IL-1β (20-fold), IL-1 receptor antagonist (2-fold) (Nold et al., 2013; Rudloff et al., 2017) type 2 cytokines (IL-4, IL-5, IL-13) (4- to 53-fold), and type 2 chemokines (CCL1, CCL17) (11- to 31-fold) (Lao et al., 2022).

      2. Five days of 65% hyperoxia

         1. Pulmonary immunity

            Transcription factors that regulate the development of all innate lymphoid cells (ILCs) (Plzf and Id2) and ILC2s (Tcf7 and Bcl11b) are up to 23% lower after hyperoxia (Lao et al., 2022). There is a 2-fold increase in type 2 helper (TH2) cells after hyperoxia (Lao et al., 2022).

         2. Pulmonary angiogenesis

            Hyperoxia elevates vascular endothelial growth factor by 1.4-fold and endothelin-1 by 1.4-fold in murine lungs (Bui et al., 2019).

      3. Fourteen days of 65% hyperoxia

         1. Alveoli

            Early changes in alveolar structure (Nold et al., 2013).

      4. Twenty-eight days of 65% hyperoxia

         1. Structural changes

            1). Alveoli

            Twenty-eight days marks the end of the alveolar stage of lung development in the mouse, similar to 2–3 years of age in humans (Backstrom et al., 2011). Hyperoxia reduces alveolar number (25%–44%), increases alveolar size (40%–230%), and reduces surface/volume ratio (18%–35%) (Figure 4) (Nold et al., 2013; Royce et al., 2016; Rudloff et al., 2017). These emphysematous changes in alveoli impair lung function by reducing the diffusing capacity of the lung (Pellegrino et al., 2005)

            
            

            
            Figure 4. Emphysematous changes in lung morphology (H&E staining) at 28 days of life after antenatal LPS and postnatal exposure to 65% hyperoxia compared to air controls in C57BL/6J mice (sourced from Rudloff et al., 2017). Scale bars 100 µm, ×200 magnification.

            
            

            2). Airways

            Hyperoxia increases airway epithelial thickness (25%), subepithelial collagen deposition (200%), expression of α-smooth muscle actin (50%), and expression of the proliferation marker cyclin D1 mRNA (260%) (Royce et al., 2016). However, hyperoxia does not alter the airway goblet cell mucus production (Royce et al., 2016).

            3). Pulmonary vascular bed

            Hyperoxia reduces the number of small vessels (4–5 µm diameter) (84%) (Bui et al., 2019; Lao et al., 2022). No differences in the expression of α-smooth muscle actin and endothelin receptor type-A after hyperoxia (Bui et al., 2019).

            4). Heart

            Hyperoxia increases cardiac fibrosis (2.2-fold) and fibrotic/inflammatory markers galectin-3 and CCL2 (C-C motif) (Bui et al., 2019).

            5). Brain

            Hyperoxia does not cause apoptosis, cortical neuronal lamination, temporal cortical width (at hippocampus), white matter myelination, hemorrhage, loss of neurons, gliosis, inflammation, or any other significant morphologic abnormalities in C57Bl/6J mice (Rudloff et al., 2017).

         2. Functional changes

            1). In vivo

            The inversely related surrogate marker time to peak velocity/right ventricular ejection time, as measured by echocardiography, increases in hyperoxic pups (0.27 compared to 0.32 in air controls) (Bui et al., 2019; Lao et al., 2022).

            2). Ex vivo

            Hyperoxia increases luminal contraction of airways to the muscarinic acetylcholine receptor agonist methacholine (10–3,000 nM) (89%) compared to air (44%) in precision cut lung slices (Royce et al., 2016). Pulmonary arterial contraction to the vasoconstrictors endothelin 1 and U46619 is not altered by hyperoxia in precision cut lung slices (Bui et al., 2019).

         3. Cellular immunity of the lung

            After hyperoxia, the overall number of immune cells (neutrophils, lymphocytes, monocytes/macrophages, natural killer cells, and dendritic cells) is approximately 3.1-fold lower. However, increased activation of macrophages and dendritic cells, as well as elevated IL-1β, is still evident in the lungs (Nold et al., 2013).

      5. Sixty days (28 days hyperoxia, followed by 32 days room air)

         Hyperoxia reduces vessel numbers at each branching generation in the murine lung (-18% in generation 2 and -21% in generation 3) (Bui et al., 2019).

         
         

6. Maintain comprehensive experimental records

   1. Record oxygen application, temperature, and humidity with power lab instrument and Chart 5 Pro software.

   2. Record assessments of animal welfare.

      
      

7. Fostering of pups (Figure 5 and Video 1)

   Adult mice are more susceptible to hyperoxia-induced oxygen toxicity than neonatal mice (Frank et al., 1978). Therefore, dams need to be rotated between room air and hyperoxia pups in a three-day cycle to limit hyperoxia-induced toxicity. It is important to conduct this procedure promptly, so that pups are in the experimental conditions for as long as possible. Top up food and water during this step to minimize unnecessary opening of the hyperoxia chamber.

   1. Remove cages from air and hyperoxia chambers.

   2. Take one pair of one hyperoxia and one air cage at a time.

      1. Swap the pups between the paired dams

         1. Air pups will be moved into the dam's cage that was previously in hyperoxia.

         2. Hyperoxia pups will be moved into the dam's cage that was previously in air.

      2. Cover the pups in bedding material from the new cage

         1. This procedure masks the scent of the other dam.

         2. This reduces the likelihood of rejection from the foster dam.

      3. Repeat for all paired cages

   3. Return cages to air and hyperoxia chambers.

      1. Pups are in the same conditions.

      2. Dams previously in hyperoxia chamber are now housed in the air chamber; however, the pups stay under their allocated experimental condition.

      3. Dams previously in air chamber are now housed in the hyperoxia chamber; however, the pups stay under their allocated experimental condition.

         
         

   
   Figure 5. Fostering of pups. Hyperoxia animals (red), air animals (blue). Air dam rotates into the hyperoxic chamber every three days. Hyperoxia dam rotates into the air chamber every three days. Pups are cross fostered every three days between dams and, as a result, they are continuously housed under the same experimental condition from day 1.

   
   
   
   Video 1. Fostering of pups

   
   

8. Subcutaneous injections of neonatal pups (Figure 6)

   1. Gently scruff the pups.

      Lift the skin above the neck.

   2. Insert the syringe at a 45° angle.

   3. Inject pups slowly.

   4. Pull out the needle slowly to prevent leakage.

      
      

      
      Figure 6. Subcutaneous injection of 11-day-old C57BL/6J pup

      
      

9. Sample preparation for lung histology and pulmonary immunity of a 28-day-old pup

   1. Anesthetize the pup with isoflurane per open-drop method (Risling et al., 2012).

   2. Euthanize the pup

      1. Cervical dislocation

         1. Place the thumb and forefinger in between the head and neck of the animal.

            Note: The spinal column should be palpable.

         2. Pinch the thumb and forefinger together and push down simultaneously until the cervical spine is cleanly separated.

            1). Ensure that you do not apply too much pressure as the trachea needs to be intact for later inflation of the lung.

            2). Pinch the foot with the forceps to ensure mouse is dead before dissecting.

   3. Dissection (Figure 7, Figure 8, and Video 2)

      1. Pin the mouse on a foam dissection board on its back with its four limbs spread out.

      2. Hook the front teeth of the mouse with a suture; then, pull tight and pin to foam board.

         This procedure helps to straighten the trachea to allow better inflation.

      3. Dissect the mouse

         1. Cut through the ventral surface of the mouse from the sternum up to the chin.

         2. Open the chest cavity such that the entire lung is visible.

         3. Expose the trachea.

            1). The trachea is enclosed by the thyroid.

            2). Grasp the thyroid with forceps and pull apart to remove.

            3). Wipe away any excess blood with gauze soaked in PBS or use a cotton tip applicator.

            4). Delicately cut away tissue around trachea so that the forceps can reach underneath.

            5). Place two strings of suture underneath the trachea.

      4. Samples for pulmonary immunity

         1. Ensure that the left lobe is clearly visible.

         2. Loop a piece of suture around the left lobe of the lung by holding the suture with your forceps and lift the lobe with a cotton tip applicator.

            Organ will stick to the cotton without being ripped or damaged. Lungs exposed to hyperoxia are fragile.

         3. Now loop the suture around the left bronchus and tie the left lobe off.

         4. Ensure knots are tight so that no leakage occurs during the right lung inflation.

         5. Cut off the left tied lobe, cut in two, and place in separate labeled 1.7 mL microtubules. Immediately snap freeze in liquid N₂ (following all safety protocols of your institution) for later mRNA and protein analysis.

            Store at -80 °C until ready for processing.

      5. Inflation of the right lung lobes

         1. Insert a 24G cannula into the trachea.

         2. Tie the cannula in place with one of the sutures threaded earlier underneath the trachea.

         3. Attach the cannula to a gravity perfusion apparatus.

         4. Infuse the right lung lobes with 4% PFA (pH 7.4) under constant 20 cm H₂O of hydrostatic pressure.

            1). Reconstitute PFA powder with PBS.

            2). Full inflation is determined by the tip of the post-caval lobe turning pointy.

         5. Tie off the trachea with the second suture in front of the cannula to prevent backflow of PFA.

         6. Place pup on ice for 10 min.

      6. Samples for histology

         1. Remove right lung lobes from thorax and place in tube with 4% PFA.

         2. Keep in 4% PFA for a minimum of 6 h.

         3. Process for paraffin embedding and sectioning.

            
            

   
   Figure 7. Sample preparation of a 28-day-old mouse. Dissect the mouse and open the chest cavity to expose lungs and trachea (A). Tie off the left bronchus and remove the left lung for later analysis of pulmonary immunity (B). Inflate the right lung lobes with 4% paraformaldehyde for histology (C).

   
   

   
   Figure 8. Pulmonary anatomy of a mouse. The left lobe (E) and the right lobes [superior lobe (A), middle lobe (B), inferior lobe (C), and post-caval lobe (D)]. At day 28, the left lobe is removed for analysis of pulmonary immunity (top third for RNA, bottom two thirds for protein). The right lobes are subsequently inflated with 4% paraformaldehyde and sections for histology are made from the inferior lobe.

   
   
   
   Video 2. Sample preparation for lung histology and pulmonary immunity of a 28-day-old mouse

Acknowledgments

The authors acknowledge the original research papers that have applied this protocol (Nold et al., 2013; Royce et al., 2016; Rudloff et al., 2017; Bui et al., 2019; Lao et al., 2022). The authors thank Ina Rudloff, Christine Bui, and Elizabeth Skuza for their considerable work with the double hit model. Investigators were supported by a co-funded Monash graduate scholarship (S.P.G.); a National Heart Foundation of Australia Future Leader Fellowship (CF14/3517), the Fielding Fellowship 2019, and NHMRC Investigator Grant 1173584 (to C.A.N.-P.); and the Hudson Institute’s Star Recruitment Fellowship, Monash University’s Larkins Fellowship, and the Fielding Foundation Fellowship 2017 (to M.F.N.). The study was furthermore supported by grants from the Australian Synchrotron (EU9230, M8925, and M9422 to C.A.N.-P., and M.F.N.), from CSL Ltd. (to C.A.N.-P. and M.F.N.), from the Jack Brockhoff Foundation (JBF3105 to C.A.N.-P., and M.F.N.), from the Rebecca L. Cooper Foundation and Perpetual’s 2016 Impact Philanthropy Program (IPAP201600780 to C.A.N.-P.), and by the Victorian Government’s Operational Infrastructure Support Program.

Competing interests

The authors declare no competing interests.

Ethics

The double hit model protocol was approved by the animal review board MMCA of Monash University.

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