Intraperitoneal Glucose Tolerance Test, Measurement of Lung Function, and Fixation of the Lung to Study the Impact of Obesity and Impaired Metabolism on Pulmonary Outcomes

Summary

The incidence of obesity is rising and increases the risk of chronic lung diseases. To establish the underlying mechanisms and preventive strategies, well-defined animal models are needed. Here, we provide three methods (glucose-tolerance-test, body plethysmography, and lung fixation) to study the effect of obesity on pulmonary outcomes in mice.

Abstract

Obesity and respiratory disorders are major health problems. Obesity is becoming an emerging epidemic with an expected number of over 1 billion obese individuals worldwide by 2030, thus representing a growing socioeconomic burden. Simultaneously, obesity-related comorbidities, including diabetes as well as heart and chronic lung diseases, are continuously on the rise. Although obesity has been associated with increased risk for asthma exacerbations, worsening of respiratory symptoms, and poor control, the functional role of obesity and perturbed metabolism in the pathogenesis of chronic lung disease is often underestimated, and underlying molecular mechanisms remain elusive. This article aims to present methods to assess the effect of obesity on metabolism, as well as lung structure and function. Here, we describe three techniques for mice studies: (1) assessment of intraperitoneal glucose tolerance (ipGTT) to analyze the effect of obesity on glucose metabolism; (2) measurement of airway resistance (Res) and respiratory system compliance (Cdyn) to analyze the effect of obesity on lung function; and (3) preparation and fixation of the lung for subsequent quantitative histological assessment. Obesity-related lung diseases are probably multifactorial, stemming from systemic inflammatory and metabolic dysregulation that potentially adversely influence lung function and the response to therapy. Therefore, a standardized methodology to study molecular mechanisms and the effect of novel treatments is essential.

Introduction

According to the World Health Organization (WHO) in 2008, more than 1.4 billion adults, aged 20 and older, were overweight with a body mass index (BMI) greater than or equal to 25; further, over 200 million men and nearly 300 million women were obese (BMI≥30)¹. Obesity and metabolic syndrome are major risk factors for a multitude of diseases. While obesity and concomitant increased white adipose tissue mass has been intimately linked to type 2 diabetes^2,3, cardio-vascular diseases including coronary heart disease (CHD), heart failure (HF), atrial fibrillation⁴ and osteoarthritis⁵, their functional roles in the pathogenesis of respiratory disorders remain poorly understood. However, epidemiological studies have demonstrated that obesity is strongly associated with chronic respiratory conditions, including exertional dyspnea, obstructive sleep apnea syndrome (OSAS), obesity hypoventilation syndrome (OHS), chronic obstructive pulmonary disease (COPD), pulmonary embolism, aspiration pneumonia and bronchial asthma^6,7,8,9. Potential mechanisms linking obesity and perturbed metabolism, e.g., insulin resistance and type II diabetes, to the pathogenesis of chronic lung disease not only comprise mechanical and physical consequences of weight gain on ventilation but also induce a chronic subacute inflammatory state^10,11. The rise of obesity and lung diseases during the last decade, coupled with the lack of effective preventive strategies and therapeutic approaches, highlights the need to investigate the molecular mechanisms to define new avenues to manage obesity-related lung diseases.

Here, we describe three standard tests, which are important basics to investigate obesity and its impact on lung structure and function in mouse models: (1) intraperitoneal glucose tolerance (ipGTT) (2) measurement of airway resistance (Res) and respiratory system compliance (Cdyn); and (3) preparation and fixation of the lung for subsequent quantitative histological assessment. The ipGTT is a robust screening test to measure glucose uptake, and thus the effect of obesity on metabolism. The simplicity of the method allows good standardization, and therefore the comparability of results between laboratories. More sophisticated methods, such as hyperglycemic clamps or studies on isolated islets, can be used for detailed analysis of the metabolic phenotype¹². Here we assess glucose tolerance to define an obesity-associated state of systemic and metabolic disorder as the basis for further studies on a pulmonary outcome. To assess the effect of obesity and metabolic disorder on lung function, we measured airway resistance (Res) and respiratory system compliance (Cdyn). To characterize lung disease, unrestrained as well as restrained methods for assessment of lung function are available. Unrestrained plethysmography in freely moving animals mimics a natural state, reflecting breathing patterns; in contrast, invasive methods, such as input impedance measurement of Res and cDyn in deeply anesthetized mice to assess dynamic lung mechanics, are more accurate¹³. Since chronic respiratory conditions are reflected by histologic alterations of the lung tissue, proper lung fixation for further analysis is imminent. The choice of the method of tissue fixation and preparation depends on the compartment of the lung which will be studied, for example, conducting airways or lung parenchyma¹⁴. Here, we describe a method that allows qualitative and quantitative assessment of the conducting airways to study the effect of obesity on asthma development.

Protocol

All animal procedures were conducted in compliance with protocols approved by local government authorities (Land NRW, AZ: 2012.A424), and were in accordance with the German animal welfare law and the regulations on the welfare of animals used for experiments or for other scientific purposes. Since lung function analysis may affect lung structure and therefore subsequent histological analyzes, the measurement of Res and Cdyn and the preparation and fixation of the lung for histomorphometry should be performed in different animals. However, measurement of Res and Cdyn following ipGTT is possible. Since stress during the ipGTT could interfere with the anesthesia needed for lung function tests, a recovery period of approximately 2 weeks after ipGTT is recommended to allow mice to recover from body weight loss and changes in blood parameters¹².

1. Preparation for Intraperitoneal Glucose Tolerance Test (ipGTT)

NOTE: After 12 h of fasting, the complete ipGTT takes approximately 2 h.

1. Since stress influences blood glucose significantly, ensure that both adaption of mice, as well as training of the scientist, are performed.
2. Transfer the animals to the experimental area under quiet and stress-free conditions.
3. Consider the application of a hypercaloric diet to induce obesity in mice. See the discussion section for further advice.
4. Fast animals for 12 h overnight, without limiting access to water. The next day, after 12 h of fasting, prepare the blood glucose meter according to the manufacturer's protocol (see table of materials) by inserting a new test strip into the test strip port.
5. Incise the tail tip using sterile scissors, while gently retaining the mouse at its tail, and immediately measure the fasted blood glucose by applying a free-flowing blood drop (minimum sample size 0.5 µL) to the test strip of the blood glucose meter.
   NOTE: A countdown timer starts on the screen after sufficient application of the blood sample. After 4 s, the test result appears on the screen.
6. Afterwards, weigh and label the animals individually using color marking.
7. Administer 2 g glucose/kg body weight via intraperitoneal injection. Ensure that the injection volume is 0.1 mL/10 g body weight (27 G needle and 1 cc syringe).
8. Subsequently, measure blood glucose after 15, 30, 60, and 120 min by applying a drop of free-flowing blood on a new test strip.
   NOTE: Blood flow can be increased by gentle massaging of the tail tip-wards. If the tail wound encrusts, clean it using a sterile swab soaked with 0.9% sodium chloride solution.
9. Allow the animals to rest in their home cages with unlimited access to water between the measurements.

2. Lung Function Analysis to Measure Res and cDyn

NOTE: For undisturbed measurement of Res and cDyn, mice need to be ventilated under deep anesthesia. Stress-free animal handling and proper monitoring of anesthesia are essential. For general instructions using sterile techniques, please review the article by Hoogstraten-Miller et al.¹⁵

1. Calibrate the plethysmograph prior to each set of experiments and prepare the study settings within the software (see Table of Materials).
2. Prior to surgery, deeply anesthetize animals via intraperitoneal injection of Xylazine (10 mg/kg body weight) and Ketamine (100 mg/kg body weight) (27 G needle and 1 cc syringe). Ensure that the injection volume is 0.1 mL/10 g per body weight.
   NOTE: Since ketamine has a proper analgesic effect in mice, no additional pain treatment is necessary. The invasive tracheal catheter/plethysmograph procedure takes approximately 5-7 minutes, then data acquisition can begin.
3. Place the mouse in the supine position on a heating pad to maintain the body temperature.
4. Cover the eyes with ointment to prevent dryness under anesthesia.
5. Constantly monitor the depth of anesthesia using the toe pinch-response.
   NOTE: Additional administration of anesthetic might be necessary to maintain a surgical plane of anesthesia.
6. Moisten the fur of the surgical area in the thyroid region with 70% ethanol.
7. Carefully incise the skin in the midline for approximately 1 cm between the jugular notch of the sternum and the tuber symphyses of the mentum by lifting it with forceps and clipping the skin under visual inspection using blunt scissors (Figure 1A).
8. Visualize the underlying subcutaneous adipose tissue and thyroid gland.
9. Expose the trachea by carefully blunt separating both thyroid lobes at the isthmus and dissection of the sternothyroid and sternothyroid muscles (Figure 1B). Be careful not to harm any vessels and cause bleeding, since this can cause adverse effects on the cardiovascular system and ultimately on the measurements.
10. Subsequently, pass a 4-0 braided surgical suture between the trachea and esophagus using blunt forceps. Carefully incise the trachea close to the larynx between the tracheal cartilages with micro scissors.
11. Intubate with a tracheal tube (0.04 inch / 1.02 mm diameter) under visual control (Figure 1C). Fix the tube via ligation with the surgical suture to avoid any leak in the system.
12. Next, move the animal to the heated bed of the body chamber and connect the tracheal tube to the face plate (Figure 1D) and turn on the ventilation by pressing the ventilation button on the front panel of the controller (Figure 1E).
13. Survey the ventilation by observing the thorax movement contemporaneously with the ventilation rate. To confirm proper placement of the tracheal tube, ensure that both sides of the thorax move simultaneously.
14. Watch the pressure signal on the computer screen (Figure 1F). Ensure that the ventilation curves are uniform. If this is not the case, detach the animal and check the surgery side. Beware of blood or mucus blocking the tracheal tube.
    NOTE: For adult animals with a body weight of 20-25 g, the ventilator settings as shown in Figure 2 are suggested in accordance with the manufacturer's recommendations.
15. To control changes in the trans-pulmonary pressure during ventilation, insert an esophageal tube (0.04 inch / 1.02 mm diameter) into the esophagus at the depth that approximates the levels of the lungs. Watch the screen while placing the tube. Place the tube where maximal pressure deflection and minimal heart artifacts can be seen on the screen.
16. After surgery, prepare the animal for the measurement. Reinject anesthesia via intraperitoneal injection of Ketamine (100 mg/kg body weight) using a 27-G needle and 1 cc syringe. Ensure that the injection volume is 0.1 mL/10 g per body weight.
    NOTE: To assess bronchial hyperreagibility, nebulize methacholine, a non-selective muscarinic receptor agonist of the parasympathetic nervous system, which induces bronchoconstriction. Data acquisition is performed in four different phases (Figure 3).
17. Start data acquisition according to the manufacturer´s protocol.
    NOTE: The software automatically guides users through the acquisition process.
18. Apply 10 µL of PBS (vehicle) on the nebulizer, and start nebulization after 5 min of acclimation. Next, follow a response phase of 3 min, where Res (cmH₂O/mL/s) and cDyn (mL/cmH₂O) are measured. At the end, provide a recovery phase of 3 min to the animal prior to the next nebulization.
19. Follow the software by stepwise application of 10 µL of increasing concentrations of methacholine (2.5 µg/10 µL, 6.25 µg/10 µL, and 12.5 µg/10 µL) on the ventilator.
20. Once all measurements have been performed and recorded, sacrifice the animal by cervical dislocation.

3. Lung Isolation for Quantitative Histomorphometric Analysis of Adult Mice

1. Deeply anesthetize the animal via intraperitoneal injection of Xylazine (10 mg/kg body weight) and Ketamine (100 mg/kg body weight) (27 G needle and 1 cc syringe). The injection volume should be 0.1 mL/10 g per body weight.
   NOTE: After reaching the state of surgical tolerance, the preparation takes approximately 5 min followed by organ perfusion and 30 min for fixation.
2. Once the animal has reached the state of surgical tolerance (negative toe pinch-response), disinfect the animal with 70% ethanol and fix the animal on a pad with surgical tape.
3. Sacrifice the animal by cardiac puncture and bleeding. Briefly, open the abdomen with a medial incision through the skin and the peritoneum using blunt scissors.
4. Locate the diaphragm head wards of the liver, and carefully separate the liver from the diaphragm.
5. Make a small incision in the diaphragm using blunt scissors, and punctate the left ventricle of the heart with a 20 G needle attached to a 2-mL syringe. Slowly exsanguinate the animal.
   NOTE: Slow and careful exsanguination is important to prevent the ventricles collapsing due to the negative pressure, inhibiting an undisturbed blood flow.
6. Dissect the lung by opening the thorax gently through a parasternal incision along the entire length of the rib cage using curved, blunt scissors.
7. Afterwards, lift the rib cage to expose the pleural cavity (Figure 3C). Remove the thymus to see the heart and lungs.
   NOTE: Optional injection of the right ventricle, followed by perfusion of the lung vascular system with ice-cold PBS and then with a fixative solution [e.g., 4% (mass/volume) paraformaldehyde (PFA)] is possible. Be aware that there is an increased risk to rupture alveolar septae and adversely affect lung structure using this method.
8. Dissect the lung by first carefully removing the heart.
9. Subsequently, pass a 4-0 braided surgical suture between the trachea and esophagus using blunt forceps.
10. Next, carefully incise the trachea close to the larynx between the tracheal cartilages, intubate with an intravenous cannula (26 G), and inflate the lung by pressure fixation at a constant pressure of 20 cm H₂O using fixative agent [e.g., 4% (mass/volume) of PFA].
11. For PFA fixation, leave the fixative for 30 min at room temperature. Afterwards, ligate the trachea and remove the cannula. Then, excise the lung carefully without harming the tissue, and store it in fixative agent at 4 °C overnight.
    NOTE: Alternatively, according to the ATS/ETS consensus paper 2.5% GA buffered OsO4, Uracil solution is used for proper tissue stabilization. For further tissue preparation, see the consensus paper by Hsia et al.¹⁴

Results

Representative results of intraperitoneal glucose tolerance test (ipGTT) (Figure 4), lung function test (Figure 5), and representative images illustrating hematoxylin and eosin stained lungs (Figure 6).

The ipGTT was performed in obese mice (blue) after 7 weeks of high-fat-diet (HFD). Standard diet-fed mice served as controls (black). Obese...

Discussion

This report provides three protocols for three different methods to analyze the impact of obesity on glucose metabolism and pulmonary outcomes. First, the glucose tolerance test offers the opportunity to analyze intracellular glucose uptake and can be indicative of insulin resistance. Second, whole body plethysmography is a technique to measure lung function and is thereby helpful to test the efficacy of novel treatments. Third, a standardized fixation protocol is essential for quantitative morphometric analysis to asses...

Materials

Name	Company	Catalog Number	Comments
GlucoMen LX	A.Menarini diagnostics, Firneze, Italy	38969	blood glucose meter
GlucoMen LX Sensor	A.Menarini diagnostics, Firneze, Italy	39765	Test stripes
Glucose 20%	B. Braun, Melsung, Germany	2356746
FinePointe Software	DSI, MC s´Hertogenbosch, Netherlands	601-1831-002
FinePointe RC Single Site Mouse Table	DSI, MC s´Hertogenbosch, Netherlands	601-1831-001
FPRC Controller	DSI, MC s´Hertogenbosch, Netherlands	601-1075-001
FPRC Aerosol Block	DSI, MC s´Hertogenbosch, Netherlands	601-1106-001
Aerogen neb head-5.2-4um	DSI, MC s´Hertogenbosch, Netherlands	601-2306-001
Forceps	FST, British Columbia, Canada	11065-07
Blunt scissors	FST, British Columbia, Canada	14105-12
Micro scissors	FST, British Columbia, Canada	15000-00
Perma-Hand 4-0	Ethicon, Puerto Rico, USA	736H	Surgical suture
Roti-Histofix 4%	Roth	P087.1	4% Paraformaldehyd
Ketaset	Zoetis, Berlin, Germany	10013389	Ketamine
Rompun 2%	Bayer, Leverkusen, Germany	770081	Xylazine