How to Measure Alterations in Alveolar Barrier Function as a Marker of Lung Injury

UNIT 24.3

Raquel Herrero1 and Gustavo Matute-Bello2 1

Hospital Universitario de Getafe, Servicio de Cuidados Intensivos, CIBER de Enfermedades Respiratorias, Getafe, Madrid, Spain 2 Medical Research Service of the Veterans Affairs Puget Sound Health Care Center and Center for Lung Biology, Division of Pulmonary and Critical Care Medicine, Department of Medicine, University of Washington, Seattle, Washington

The alveolar capillary membrane maintains the proper water and solute content of the epithelial lining fluid at the alveolar air-liquid interface, which is critical for adequate gas exchange in the lung. This is possible due to the alveolar fluid clearance (AFC) capacity of this membrane that assists in the removal of salt and water from the alveolar air spaces. The alveolar capillary membrane also provides a barrier that restricts the passage of proteins and water from the interstitial and vascular compartments into the alveolar air spaces. This restricted passage is due to the presence of tight junctions between adjacent alveolar epithelial cells. Severe injury to the alveolar epithelial/endothelial membrane results in increased protein permeability and impairment of AFC, which leads to the formation of protein-rich edema with the consequent deterioration of gas exchange. Many animal models of lung injury, focused on damage of the alveolar-capillary membrane, assess the AFC capacity and the barrier function. We describe a simple method to assess the AFC rate in normal and pathological conditions in mice. We also describe two complementary methods to assess the alveolar-capillary barrier function, which require measuring the concentration of endogenous plasma proteins in bronchoalveolar lavage fluid and detection C 2015 by of tight-junction proteins in lung tissue by immunofluorescence.  John Wiley & Sons, Inc. Keywords: lung injury r alveolar fluid clearance r barrier function r protein permeability r tight-junction proteins r mice

How to cite this article: Herrero, R. and Matute-Bello, G. 2015. How to Measure Alterations in Alveolar Barrier Function as a Marker of Lung Injury. Curr. Protoc. Toxicol. 63:24.3.1-24.3.15. doi: 10.1002/0471140856.tx2403s63

INTRODUCTION In normal lungs, the alveolar epithelial membrane provides a barrier function that limits the movement of liquid and proteins from the interstitial and vascular spaces to the air spaces. The alveolar epithelial membrane is also responsible for removing salt and water from the distal air spaces of the lung, which is known as the alveolar fluid clearance (AFC) capacity. Both the barrier and the AFC functions contribute to maintain the proper alveolar air-liquid interface, which is critical for adequate gas exchange and host defense against viral and bacterial pathogens (Matthay et al., 2005). Experimental animal studies show that substantial injury to the alveolar epithelial/endothelial membrane results in increased protein permeability and impairment of AFC. These alterations lead to alveolar flooding with high molecular weight proteins, Current Protocols in Toxicology 24.3.1-24.3.15, February 2015 Published online February 2015 in Wiley Online Library (wileyonlinelibrary.com). doi: 10.1002/0471140856.tx2403s63 C 2015 John Wiley & Sons, Inc. Copyright 

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disordered repair and, consequently, worsening of gas exchange (Matthay et al., 2005). Increased protein permeability and alteration of the AFC capacity occur in most patients with acute respiratory distress syndrome (ARDS), which is also characterized by important alveolar epithelial and endothelial cell damage; importantly, the alteration of the AFC has been associated with increased mortality (Ware and Matthay, 2001). Many of the various methods to assess the function of the alveolar-capillary membrane in different animal models of lung injury focus on assessing the AFC capacity and the barrier function. Prior studies from different groups demonstrated that the rate of shortterm AFC is not adversely affected by the absence of perfusion or ventilation to the lung (Effros et al., 1987; Sakuma et al., 1993, 1997; Carter et al., 1996). Therefore, methods to quantify the isosmolar AFC over a short period of time (30 min or 60 min) have been developed that have the advantage of being performed in the whole animal without the need of lung perfusion or ventilation (Garat et al., 1998). There are also several methods to evaluate the alveolar-capillary barrier function, for example by assessing lung protein permeability using bronchoalveolar lavage (BAL) samples (Herrero et al., 2013). Evaluation of tight-junction proteins may provide further evidence of the disruption of the alveolar epithelial barrier and the consequent increased protein permeability in the lung (Mazzon and Cuzzocrea, 2007). We describe a useful and easy method to assess the AFC rate in mice with normal and pathological conditions as well as after pharmacological stimulation. We further describe two complementary methods to assess the alveolar-epithelial barrier function, which require measuring the concentration of endogenous plasma proteins in BAL fluid and evaluating the expression of tight-junction proteins in lung tissue by immunofluorescence. BASIC PROTOCOL 1

MEASUREMENT OF ALVEOLAR FLUID CLEARANCE (AFC) IN SITU IN MOUSE LUNGS The main aim of this protocol is to measure the AFC as an indicator of sodium and water transport across the distal and alveolar lung spaces in mouse lungs in situ. This technique is performed in mice that are previously sacrificed and have no blood flow in the lungs, which allows one to exclusively determine the capacity of water absorption of the distal and alveolar epithelial/endothelial cells (without the circulatory component). The measurement is performed in the whole animal, without extracting the lungs from the thoracic cavity. We first describe the preparation of an instillate that contains a known concentration of fluorescently-labeled albumin that will be instilled intratracheally into the lung. We then describe the instillation procedure, the timing of measurements, and a simple equation to calculate the AFC.

Materials 8 week-old mice (body weight: 25 to 30 g) Human serum albumin (stock 50 mg/ml; Baxter) Ringer’s lactate solution (Baxter) Fluorescein isothiocyanate (FITC)-tagged human serum albumin (Sigma-Aldrich) 7% sodium chloride solution (see recipe) Pentobarbital (Nembutal sodium; Abbot Laboratories) Alcohol

Assessment of AlveolarCapillary Membrane Dysfunction

Osmometer 18-G Terumo Surflo ETFE i.v. catheter (Terumo Corporation) Rectal temperature probe Thermostatically controlled pad Infrared lamp (Thermo Fisher Scientific)

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Mouse dissection kit (including dissection board, gauze, tweezers, scissors, and curved sharp-tip small scissors) 2-0 silk suture thread Balance (to weigh mice) 1-ml syringes 25-G needle P200 pipet For continuous positive airway pressure and 100% oxygen flow: Oxygen regulator PVC tubing (Value Plastics) T-tube fittings with 200 series barbs (Value Plastics) 3-way stopcocks with luer connections and male lock (Value Plastics) 96-well plate (white/clear bottom, TC surface; BD Falcon) Fluorescence spectrophotometer (Thermo Fisher Scientific) Centrifuge NOTE: Avoid lipopolysaccharide (LPS) and microbial contamination throughout the instillation procedure. Perform all instillations inside a biosafety hood. Preparation of instillate buffer for AFC measurement 1. Prepare solution of 5% human serum albumin in Ringer’s lactate solution. 2. Add FITC-tagged human serum albumin (used as an alveolar protein tracer) for a final concentration in the instillate fluid of 0.16 mg/ml and gently mix the solution. 3. Measure osmolality of the instillate by using an osmometer, and adjust it to 324 mOsm/kg by adding small drops of 7% sodium chloride solution (add 5 μl at a time) in order to make the instillate fluid isosmolar to plasma before instillation. This instillate fluid must be prepared immediately before use.

Intratracheal instillation 4. Weigh mice. 5. Euthanize mice with an intraperitoneal injection of pentobarbital (120 mg/kg) using a 1-ml syringe equipped with a 25-G needle. 6. Place mice in a decubitus position throughout the experiment. 7. Within 2 min of death, make a middle incision in the neck skin and incise the muscle longitudinally to expose the trachea, using scissors. Place a 2-0 silk suture thread under the trachea. 8. Make a small incision on the frontal face of the trachea by using the tip of a curved sharp-tip small scissor. Cannulate trachea through the incision with an 18-G trimmed i.v. catheter (Terumo Surflo ETFE i.v. catheter). Take care not to cut through the trachea.

Secure the trachea and its catheter with a 2-0 silk suture thread around the trachea. 9. Monitor body temperature continuously by using a rectal temperature probe and maintain it at 37o to 38°C with a thermostatically controlled pad and additional light lamps, if needed. The lamp is cycled on and off to maintain the core temperature at 37o to 38°C. Alternatively, body temperature may also be monitored by introducing a temperature probe (Yellow Springs Instrument [YSI]) into the abdominal cavity via a 0.5-cm incision.

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Figure 24.3.1 Application of a continuous positive airway pressure (CPAP) at 5 cm H2 O with 100% oxygen flow into mouse lungs through an intratracheal catheter. 1) Oxygen regulator 2) PVC tubing 3) T-tube fitting with 200 series barbs. 4) Stopcocks with luer connections; 3-way; male lock. 5) Free luer of stopcock used for lung fluid instillation and aspiration. 6) Intratracheal catheter. 7) Mouse. 8) Water column with a tube submerged under 5 cm of water. 9) Heating pad.

10. Apply continuous positive airway pressure (CPAP) at 5 cm H2 O with 100% oxygen through the intratracheal catheter (see Fig. 24.3.1). CPAP and oxygen are applied throughout the experiment to prevent airway and alveolar collapse, maintain a homogeneous distribution of the instillate, and ensure adequate tissue oxygenation.

11. Load 1-ml syringe with a volume of 12 ml/g of body weight of the instillate solution and place the 1-ml syringe in the free port of the stopcock that is connected to the tracheal cannula (Fig. 24.3.1). 12. Once mice are normothermic and well connected to the CPAP/oxygen (O2 ) tubing system (Fig. 24.3.1), instill desired fluid volume into the tracheal cannula to both lungs through the stopcock over 30 sec then insufflate 0.1 ml of room air with a 1-ml syringe into the tracheal catheter to clear the catheter dead space and position the fluid in the alveolar spaces. Make sure to close the stopcock luer connected to the CPAP/O2 tubing system during instillation.

13. Close free stopcock luer to restore the CPAP and O2 flow administration into the lungs. 14. After 1 min or 30 min, aspirate lung fluid gently through the tracheal cannula by pipetting out (P200-pipet) while applying a gentle compression on the mouse chest to aid in lung fluid to release.

Assessment of AlveolarCapillary Membrane Dysfunction

15. Centrifuge the aspirated fluid at 300 × g for 10 min, and take the supernatant. Transfer samples of the aspirated fluid supernatant to a 96-well plate and measure the fluorescence intensity of the samples by spectrophotometry at an excitation wavelength of 495 nm (485/20), and an emission wavelength of 530 nm (530/25).

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16. Calculate concentration of FITC albumin of the aspirated fluid samples from the fluorescence measurements based on a previously established lineal relationship between different concentrations of FITC albumin and its fluorescence. At the end of the experiment, remove both lungs through a median sternotomy and flash freeze them in liquid nitrogen for further determinations (e.g., FITC albumin that remains in the lung) if needed.

Calculation of the AFC The alveolar fluid collected at 1 min is considered the initial total albumin concentration of the alveolar fluid in the mice. The difference in the intensity of fluorescence in the lung fluid at 30 min with respect to 1 min is used to calculate the amount of water that is cleared from the airspace. 17. Calculate the AFC by measuring the increase in the concentration of FITC albumin between the 1-min and 30-min times using the following relationship, as previously described (Smedira et al., 1991; Yue and Matalon, 1997): AFC = (1 − C1 min /C30 min )/0.95 where C1 min and C30 min are the FITC albumin concentration of the alveolar samples at 1 min and 30 min after instillation, respectively.

PROTEIN PERMEABILITY OF THE ALVEOLAR-CAPILLARY MEMBRANE IN MICE

BASIC PROTOCOL 2

The selective barrier function of the alveolar-capillary membrane is lost in severely injured lungs. This damage is manifested by the leakage of protein-rich fluid from the vascular circulation into the interstitial and/or alveolar spaces. Many of the methods that assess this alteration require the measurement of different tracers in the bronchoalveolar lavage (BAL) fluid. Here we will describe a simple method to obtain BAL fluid in mice and list the most commonly endogenous markers that can be measured in BAL fluid and plasma in order to determine changes in the permeability of the alveolar-capillary membrane.

Materials 8 week-old mice (body weight: 25 to 30 g) Pentobarbital (Nembutal sodium; Abbot Laboratories) Heparin Bronchoalveolar lavage fluid buffer (see recipe) ELISA kit (IgM, α2 -macroglobulin; bicinchoninic acid method [BCA assay]) Mouse dissection kit (including dissection board, gauze, tweezers, scissors, and curved sharp-tip small scissors) 2-0 silk suture thread Balance (to weigh mice) 1-ml syringes equipped with 25-G needles 23-G sterile needles (if sampling blood) 18-G Terumo Surflo ETFE i.v. catheter (Terumo Corporation) 5-ml polypropylene tubes Centrifuge Thermostatically controlled pad Respiratory Toxicology

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Bronchoalveolar lavage 1. Weigh mice before euthanasia. 2. Euthanize mice with an intraperitoneal injection of pentobarbital (120 mg/kg) using a 1-ml syringe equipped with a 25-G needle. Keep animals in a cage with bedding placed on a heating pad in order to prevent the development of hypothermia while being anesthetized.

3. Once mouse is properly sedated, place it in a horizontal position over a heating pad. 4. Exsanguinate mouse by direct cardiac puncture using a 1-ml syringe and a 23-G needle with a trace amount of heparin. 5. Open thoracic cavity and make a small midline incision in the neck skin on the trachea and allow the lungs to collapse spontaneously. Separate salivary glands and incise muscle longitudinally to expose the trachea. Place a 2-0 silk suture thread under the trachea. 6. Make a small incision on the frontal face of the trachea by using the tip of a curved sharp-tip small scissor. Cannulate trachea through the incision with an 18-G trimmed i.v. catheter (Terumo Surflo ETFE i.v. catheter). Take care not to cut through the trachea.

7. Secure trachea and its catheter with a 2-0 silk suture thread around the trachea.

Collection of BAL fluid from both lungs 8. Instill 1 ml bronchoalveolar lavage fluid buffer (saline/EDTA buffer; pre-warmed to 37°C) into the tracheal cannula to both lungs using a 1-ml syringe and wait 30 sec. Do not remove syringe from the tracheal cannula.

Then, slowly aspirate lung fluid back with the same syringe. Remove syringe from the tracheal cannula and inject the recovered lavage fluid into a 5-ml tube placed on ice. Repeat the procedure three more times per mouse. Pool all aliquots together and place on ice.

Collection of BAL fluid from one lung 9. Clamp and suture hilum of the lung (that we want to exclude from the lavage) with a 2-0 silk suture thread. Instill 0.5 ml saline/EDTA buffer (pre-warmed at 37o C) with a 1-ml syringe into the tracheal cannula to the lung and wait 30 sec. Do not remove syringe from the tracheal cannula.

Then, slowly aspirate lung fluid back with the same syringe. Remove syringe from the tracheal cannula and inject recovered lavage fluid into a 5-ml tube placed on ice. Repeat the procedure three more times per mouse. Pool all aliquots together and place on ice. The other lung can be flash frozen or fixed for other assessments.

10. Centrifuge BAL fluid at 300 × g for 15 min, remove supernatant aseptically, and store it in individual aliquots at -80°C. Keep the cell pellets on ice (maximum 6 hr) for cell analysis if needed. Assessment of AlveolarCapillary Membrane Dysfunction

Measurements of alteration of the alveolar-capillary barrier 11. Measure total protein concentration in the BAL fluid by bicinchoninic acid (BCA) method.

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The BCA method is a simple and quick method to measure total protein concentrations.

12. Measure concentration of high molecular weight plasma proteins such as IgM and/or α2 -macroglobulin in BAL fluid by ELISA (Herrero et al., 2013).

EVALUATION OF EXPRESSION OF TIGHT-JUNCTION PROTEINS IN MOUSE LUNGS BY IMMUNOFLUORESCENCE

BASIC PROTOCOL 3

The presence of tight-junction (TJ) proteins between adjacent alveolar epithelial cells is responsible for the restricted passage of water and macromolecules across the alveolar epithelium into the alveolar spaces. The TJ barrier is composed of transmembrane proteins (occludin, claudins) that interact at the membrane with scaffolding proteins (zonula ocludens). The cellular distribution of TJ proteins in both frozen and fixed tissues may be assessed by immunofluorescence (IF) techniques. Here, we describe a method of indirect IF in formalin-fixed and paraffin-embedded lung tissue, including an antigen retrieval step necessary for antigen detection in over-fixed tissue. In this method, the primary antibody that specifically recognizes the TJ protein of interest is unlabeled and the second anti-immunoglobulin antibody directed toward the primary antibody is tagged with a fluorescent dye. The fluorescence can then be visualized using fluorescence or confocal microscopy and quantified using image-processing software.

Materials Paraffin-embedded murine lung tissue sections (4-μm thick) Xylene Ethanol (100%, 95%, 70%) 1× Phosphate-buffered saline (PBS; see recipe) Distilled H2 O Citrate buffer, pH 6.0 (see recipe) Proteinase K working solution (see recipe) Phosphate-buffered saline/Tween (PBST; see recipe) Blocking buffer (5% bovine serum albumin [BSA] in PBST, freshly prepared) Antibody diluent buffer (1% bovine serum albumin in PBS) Primary antibody against the protein of interest (diluted at an appropriate concentration in antibody diluent buffer) Secondary antibody against host species of primary antibody, conjugated to a fluorescent dye (e.g., fluorescein, Alexa Fluor, Texas Red; diluted at an appropriate concentration in 1× PBS) DAPI nucleic acid stain (stored at 5 mg/ml; working concentration: 0.5 μg/ml; Invitrogen) Fluorescent mounting medium (Dako) Clear nail polish (with applicator) Antifade reagent (optional) Oven or slide warmer Pressure cooker Plastic Coplin jar, with slide racks Liquid repellent slide marker pen Cover slips 50 × 22 mm (No. 1.5; Fisherbrand cat. no. 12-544-D; Fisher Scientific) Humidified chamber Paper towels Kimwipes (Fisher Scientific) Fluorescence microscope NOTE: Fixed tissue should be embedded in paraffin and sectioned onto slides. Do not allow slides to dry at any time during the whole procedure.

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Deparaffinize/hydrate sections 1. Heat slides in the oven or slide warmer for 20 to 40 min at 60°C. 2. Deparaffinize sections in two changes of xylene for 5 min each. 3. Incubate sections in two changes of 100%, 95%, and 70% ethanol for 5 min each. 4. Rinse twice in distilled water for 5 min each.

First antigen retrieval step 5. Bring tap water to boil in a pressure cooker with the lid open. 6. Immerse slides in a plastic Coplin jar or slide rack containing just enough citrate buffer (pH 6.0) to cover the slides, and cover with lid. 7. As soon as the pot water starts boiling, immerse Coplin jar containing the slides into the boiling water in the pressure cooker. Make sure that the water in the pot covers two thirds of the Coplin jar.

8. Seal and lock pressure cooker and maintain high heat to build pressure. When the indicator of the pot reads maximal pressure, turn heat down to low and wait 3 more min. 9. Remove pot from heat. Place pot under cold running water to cool it down and release pressure quickly before unsealing the lid. Remove Coplin jar and let it cool down on the bench top for 20 min at room temperature. Wash sections in PBS three times for 5 min each.

Second antigen retrieval step 10. Outline tissue sections using a liquid repellent slide marker pen. 11. Cover tissue sections with proteinase K working solution and incubate 10 to 30 min at 37°C in a moist atmosphere. Optimal incubation time may vary depending on tissue type and degree of fixation, and should be determined previously.

12. Wash sections in PBS three times for 5 min each.

Blocking 13. Remove slides from rack, tap off excess PBS onto paper towel and lay flat in humidified chamber. 14. Add enough blocking buffer to cover each tissue section, using a pipet. 15. Close off humidified chamber and incubate 1 hr at room temperature. 16. Wash slides three times in PBS for 5 min each.

Primary antibody 17. Tap off excess PBS onto paper towel and lay flat in humidified chamber. 18. Add primary antibody at appropriate dilution in primary antibody dilution buffer to each section. Incubate overnight at 4°C in a humidified chamber. 19. Wash slides in PBS three times for 5 min each. Assessment of AlveolarCapillary Membrane Dysfunction

Secondary antibody 20. Tap off excess PBS onto paper towel and lay flat in humidified chamber. 21. Add secondary antibody diluted in PBS to each section.

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22. Close humidified chamber and incubate 1 to 2 hr at room temperature in the dark.

DAPI stain 23. Wash slides in PBS three times for 5 min each. 24. Tap off excess PBS onto paper towel and lay flat on paper towel. 25. Add DAPI at working concentration to each section. 26. Protect from light and incubate 10 min at room temperature. 27. Wash sections in PBS three times for 5 min each.

Mount and cover slip 28. Remove one slide at a time from PBS and tap off excess onto paper towel. Lay flat on a paper towel and remove any remaining PBS outside the tissue sections with a folded Kimwipe. 29. Add 3 drops fluorescence mounting medium to the slide. Cover slides with cover slips and dry 1 hr in the dark at room temperature. 30. Seal edges of cover slip with nail polish and dry15 min in the dark. Store slides at 4°C in the dark.

Analysis 31. Analyze slides under fluorescence microscopy. Fluorescence excitation/emission maxima for DAPI: 358/461 nm. REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.

Bronchoalveolar lavage fluid buffer Dissolve 9 g of sodium chloride in 1 liter of water plus 0.6 mM EDTA Store at 4°C for up to 6 months. Citrate buffer, pH 6.0 Add trisodium citrate dihydrate (0.29 g) to 100 ml distilled H2 O. Mix to dissolve. Adjust pH to 6.0. Add Triton X-100 (100 μl). Mix well. Store at room temperature for up to 1 hr. Phosphate-buffered saline (PBS) For a 10× solution, per liter: 80 g NaCl 2 g KCl 14.4 g Na2 HPO4 2.4 g KH2 PO4 1 liter distilled H2 O Mix to dissolve. Adjust pH to 7.4 then autoclave. Dilute 1:10 before use. Store PBS at room temperature for up to 6 months. Phosphate-buffered saline/Tween (PBST) Add 0.3% Triton X-100 (v/v) to PBS. Stir to dissolve. Store at room temperature for up to 6 months. Respiratory Toxicology

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Proteinase K working solution, (1×, 20 μg/ml) 1 ml of Proteinase K Stock Solution (20×, 400 μg/ml) 19 ml of TE Buffer (10 mM Tris-Cl, pH 7.5; 1 mM EDTA) Mix well. Aliquot and store at 4°C for 1 month. Prewarm to 37°C before use. Sodium chloride solution, 7% 7 g NaCl 100 ml distilled H2 O Mix to dissolve. Autoclave. Store at room temperature for up to 6 months. COMMENTARY Background Information

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The present method to measure the AFC capacity has been shown to be reliable under both normal (Jayr et al., 1994; Garat et al., 1998) and pathological conditions in different animal models of lung injury (Garat et al., 1995, 1998; Pittet et al., 1996; Rezaiguia et al., 1997; Herrero et al., 2013). Murine models allow the study of the function of the alveolarcapillary membrane in normal and injured lungs with some advantages over other animals, such as faster basal clearance rates in non-perfused lungs compared with those reported in similar in situ experimental models in other species (Sakuma et al., 1993, 1997; Garat et al., 1998). Also, it was suggested that the stimulated clearance rate in mice is similar to the fast rates of AFC in patients recovering from hydrostatic pulmonary edema (Garat et al., 1998). Bronchoalveolar lavage is a useful technique that allows the study of different components of the alveolar epithelial fluid, including the concentration of a variety of proteins and other molecules, and the influx of inflammatory cells to the air space (Herrero et al., 2011, 2013). Different measurements can be performed in BAL fluid to determine protein permeability. These include the concentration of endogenous proteins such as total protein, albumin and high molecular weight plasma proteins (IgM or α2 -macroglobulin; Lipke et al., 2010; Herrero et al., 2013). Also, protein permeability can be assessed by determining the leakage of exogenous high molecular weight labeled molecules (125 I-albumin or fluorescent high molecular weight dextran) or of Evans blue dye into the alveolar space (Garat et al., 1995; Modelska et al., 1999; Xie et al., 2013). The alteration of protein permeability may be supported by evidence of a disruption of tight-junction proteins by different immunohistochemical techniques (Mazzon

and Cuzzocrea, 2007; Xie et al., 2013). Immunofluorescence (IF) is a common laboratory technique in which antibodies are chemically conjugated to fluorescent dyes such as fluorescein isothiocyanate (FITC), tetramethyl rhodamine isothiocyanate (TRITC), Alexa Fluor or others. This technique is a powerful tool for determining the cellular distribution of antigens in both frozen and fixed tissues. IF has advantages over the standard immunohistochemistry of combining high sensitivity with high resolution in the visualization of antigens in the tissues.

Critical Parameters and Troubleshooting The AFC measurement in this in situ mouse model and the BAL fluid for the protein permeability quantification are simple procedures and are easy to perform, but require certain training and surgical skills. Therefore, these methods should be performed by the same investigator, and always follow the same protocol to achieve adequate quality and reproducibility. In the AFC and BAL fluid protocols, it is important to preferentially use a flexible catheter rather than a needle for the tracheal cannulation. This will minimize the risk of accidental rupture of the trachea and will facilitate the retrieval of the airspace fluid. In the AFC, in particular, the administration of the instillate to the tracheal cannula through a stopcock will minimize the direct manipulation of the cannula and the disturbance of the CPAP and O2 flow. Body temperature is a critical factor that must be strictly controlled in all animal models. This is particularly important in the measurement of the AFC because low body temperature is likely to decrease the basal AFC capacity in mice. Also, LPS or live microorganism contamination of the instillate fluid can also modify the AFC results, as they have been

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Figure 24.3.2 AFC in basal conditions and under the effect of isoproterenol and amiloride in mice. Effect of β2 -adrenergic stimulation mediated by isoproterenol (1 × 10-5 M) and Na+ channel blockage mediated by amiloride (1 × 10-3 M) on the alveolar fluid clearance in mouse lungs. Both isoproterenol and amiloride were added to instillate fluid containing FITC albumin. After 1 min or 30 min, lung fluid was aspirated and read in a fluorescence reader, and the AFC was calculated for both time points. Each dot represents an individual mouse. Horizontal bars represent medians. (*) P < 0.05 vs all other groups.

shown to increase AFC in mouse lungs (Garat et al., 1995; Rezaiguia et al., 1997). Therefore, it is important to maximize the sterile conditions during the protocol as much as possible, and check for LPS contamination in the instillate fluid. For the AFC measurement in mice, previous investigators have used different instillate volumes ranging from 1.5 to 20 ml/kg body weight (Matthay et al., 1982; Basset et al., 1987; Garat et al., 1998). In our experiments, we found that 12 ml/kg was sufficient to retrieve enough air-space sample to measure FITC albumin and total protein at the end of the 30-min experiment (Fig. 24.3.2). If your experiment lasts more than 30 min, larger instillate volumes, closer to 20 ml/kg body weight, will facilitate sampling of alveolar fluid at the end of the experiment. According to previous studies, the fluid clearance rate is slightly faster when using the lower instillate volume, and the basal clearance rate may be decreased when using the larger instillate volume (Garat et al., 1998). Therefore, we do not recommend instilling volumes larger than 20 ml/kg (in general, no more than 0.5 ml). In the bronchoalveolar lavage technique, the lavage needs to be performed several times to maximize air space fluid recovery. To prevent destruction of the lung structure, the total volume of lavage should not exceed 1 ml

and there should be no more than four lavages when using this volume. Formalin or other aldehyde fixation in lung tissue forms protein cross-links that may mask the antigenic sites in tissue specimens, which can result in weak or false negative staining for detection of the targeted proteins by IF. To optimize the fluorescence signal when the tissue is over fixed, it is important to perform two antigen retrieval procedures: 1) citrate buffer at high temperature and high pressure atmosphere, and 2) proteinase K digestion. Both procedures cause breakdown of the protein cross-links and help unmask the antigens and epitopes in formalin-fixed and paraffinembedded tissue sections, thus enhancing the staining intensity of antibodies. Nevertheless, this antigen retrieval method may cause tissue damage in under-fixed tissues. Therefore, it is always important to select an appropriate incubation time and temperature (20° to 60°C) for the antigen retrieval procedure in order to avoid over-digestion of the tissues. Other factors that determine the success of IF include the specificity of the antibodies, the specimen preparation and the fluorescence detection instrument. All IF experiments require the correct negative control to assess the specificity of the staining. Photobleaching and quenching represent the main limitations of fluorescence microscopy. Photobleaching is the photochemical destruction of a fluorophore under highintensity illumination, and quenching is the process of decreasing the intensity of the fluorescence emission that occurs most frequently by processes such as temperature, high oxygen concentrations, and self-aggregation of dyes. The most general recommendation to reduce photobleaching and quenching is to keep the tissue in a cool and dark environment (for example, in a refrigerator at 4o C). Also, photobleaching and quenching can be minimized by: 1) using fluorophores with high quantum efficiency (Alexa Fluor dyes are a good option), 2) decreasing the excitation light in both intensity (lamp intensity) and duration (for example, using shutter control to minimize light exposure time), coupled with the use of a low-light level digital camera designed specifically for fluorescence microscopy, and 3) using oxygen scavengers and other specific antifade reagents that are commercially available. Autofluorescence is the fluorescence background that originates from the tissue and represents another artifact that compromises the detection of the fluorescence signal. Fixation with aldehydes, particularly glutaraldehyde,

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Figure 24.3.3 Total protein and IgM concentrations in BAL fluid after lung injury induced by a proapoptotic agent in mice. Effect of a pro-apoptotic agent (recombinant sFasL) on the concentration of total protein (A) and IgM (a plasma protein of large size, 900 kDa; B) in BAL fluid. Mice were treated with intratracheal instillation of recombinant human sFasL, then studied 16 hr later. As a control, mice were treated with PBS via intratracheal instillation. Each dot represents an individual mouse. Horizontal bars represent medians. (*) P< 0.05 vs PBS.

Figure 24.3.4 Immunofluorescence for ZO-1 detection in mouse lungs. Paraffin-embedded sections of mouse lungs. (A-B) section stained with a primary antibody to ZO-1 and Alexa Fluor 546-conjugated secondary antibody. Stains were visualized by fluorescence microscopy using red wavelength (ZO1-1) or blue wavelength (DAPI). (C-D) negative control section incubated with Alexa Fluor 546-conjugated secondary antibody but without primary antibody. Original magnification 200×.

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can result in high levels of autofluorescence. This can be minimized by selecting optical filters and probes that absorb and emit at longer wavelengths that maximize the fluorescence signal relative to the autofluorescence (e.g., fluorescent dye that can be excited at >500 nm). For the IF technique, it is mandatory to control for the autofluorescence by re-

placing primary antibody with blocking buffer in some control sections.

Anticipated Results The impact of injury stimuli on the AFC capacity and the alveolar epithelial/endothelial barrier function varies depending on the type and severity of lung injury and the time

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Table 24.3.1 Timeline for Procedures

Step(s)

Time required

Basic Protocol 1. Measurement of alveolar fluid clearance Preparation instillate fluid

10 min

Tracheal cannulation Intratracheal instillation Lung fluid aspiration

40 min

Centrifuge lung fluid Measurement of FITC-albumin concentration in lung fluid

20 min

Basic Protocol 2. Protein permeability of the alveolar-capillary membrane Bronchoalveolar lavage and centrifugation of fluid

30 min

Basic Protocol 3. Detection of tight-junction proteins in lungs by immunofluorescence Deparaffinize/hydrate sections

85 min

First antigen retrieval step: citrate buffer

60 min

Second antigen retrieval: proteinase K

55 min

Blocking

75 min

Primary antibody

Overnight

Secondary antibody

60-120 min

DAPI

25 min

Mount, cover and sealing

75 min

after the insult. In the present protocol, AFC is measured by the progressive increase in the protein concentration of a 5% albumin solution instilled into the distal air spaces of the lungs over 30 min. Using the present method, the basal AFC in normal mouse lungs is 30% ± 7.7 of instilled fluid (Fig. 24.3.2). This murine model is useful to investigators in the field of alveolar epithelial fluid transport and its regulation. The activity of Na+ -channels and Na+ -K+ -ATPase can be upregulated via β-adrenergic stimulation (Suzuki et al., 1995; Matalon et al., 1996; Sartori and Matthay, 2002). In mice, up-regulation of AFC mediated by β-adrenergic stimulation is dependent on β1 -receptors (Matthay et al., 2005; Mutlu and Sznajder, 2005). Isoproterenol (a βadrenergic agonist) has been tested in mouse lungs in normal and pathological conditions (Garty and Palmer, 1997; Farnand et al., 2011; Herrero et al., 2013). In our previous studies, the magnitude of stimulation with isoproterenol for 30 min is modest (20% over basal levels), perhaps partly because basal clearance is already high in mice (Fig. 24.3.2). This model also allows quantification of the changes in AFC capacity in the presence of moderately severe lung injury (Herrero et al., 2013).

In the bronchoalveolar lavage technique, each instillation volume of 1 ml results in good expansion of the lungs and facilitates the distribution of the instillate fluid to the distal airway and alveolar air spaces. This permits the instillate fluid to mix with the endogenous alveolar fluid and facilitates a good recovery of this mixture from the alveolar fluid (Fig. 24.3.3). All these considerations are particularly important when the lung is severely injured and there are bronchial obstructions and alveolar collapse. Immunofluorescence (IF) is a very useful laboratory technique for determining the cellular distribution of different antigen. IF is more frequently used in frozen tissue because of the difficulties in antigen detection in fixed tissues, as mentioned above. The present IF technique has been develop to detect antigens using fluorescent dyes even in over-fixed paraffinembedded lung tissue (Fig. 24.3.4).

Time Considerations Table 24.3.1 summarizes the timelines associated with all the protocols. Note that several steps are very time consuming.

Acknowledgement The authors thank Venus A. Wong (Medical Research Service of the Department of

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Veterans Affairs, Seattle, Washington) for her technical assistance with animal protocols, and Mar Granados (Pathology Service of the Hospital Universitario de Getafe) for her technical assistance with the immunofluorescence. This work was supported in part by grants from the Medical Research Service of the Department of Veterans Affairs and by HL-081764, HL-083044, HL-075381, and P30 DK-17047 from the National Institutes of Health, and PI12/02451 from Instituto de Salud Carlos IIIMinisterio de Economia y Competitividad.

Literature Cited Basset, G., Crone, C., and Saumon, G. 1987. Fluid absorption by rat lung in situ: Pathways for sodium entry in the luminal membrane of alveolar epithelium. J. Physiol. 384:325-345. Carter, E.P., Matthay, M.A., Farinas, J., and Verkman, A.S. 1996. Transalveolar osmotic and diffusional water permeability in intact mouse lung measured by a novel surface fluorescence method. J. Gen. Physiol. 108:133-142. Effros, R.M., Mason, G.R., Sietsema, K., Silverman, P., and Hukkanen, J. 1987. Fluid reabsorption and glucose consumption in edematous rat lungs. Circ. Res. 60:708-719. Farnand, A.W., Eastman, A.J., Herrero, R., Hanson, J.F., Mongovin, S., Altemeier, W.A., and Matute-Bello, G. 2011. Fas activation in alveolar epithelial cells induces KC (CXCL1) release by a MyD88-dependent mechanism. Am. J. Respir. Cell Mol. Biol. 45:650-658. Garat, C., Carter, E.P., and Matthay, M.A. 1998. New in situ mouse model to quantify alveolar epithelial fluid clearance. J. Appl. Physiol. 84:1763-1767. Garat, C., Rezaiguia, S., Meignan, M., D’Ortho, M.P., Harf, A., Matthay, M.A., and Jayr, C. 1995. Alveolar endotoxin increases alveolar liquid clearance in rats. J. Appl. Physiol. 79:20212028. Garty, H. and Palmer, L.G. 1997. Epithelial sodium channels: Function, structure, and regulation. Physiol. Rev. 77:359-396. Herrero, R., Tanino, M., Smith, L.S., Kajikawa, O., Wong, V.A., Mongovin, S., Matute-Bello, G., and Martin, T.R. 2013. The Fas/FasL pathway impairs the alveolar fluid clearance in mouse lungs. Am. J. Physiol. Lung Cell Mol. Physiol. 305:L377-L388. Herrero, R., Kajikawa, O., Matute-Bello, G., Wang, Y., Hagimoto, N., Mongovin, S., Wong, V., Park, D.R., Brot, N., Heinecke, J.W., Rosen, H., Goodman, R.B., Fu, X., and Martin, T.R. 2011. The biological activity of FasL in human and mouse lungs is determined by the structure of its stalk region. J. Clin. Invest. 121:1174-1190. Assessment of AlveolarCapillary Membrane Dysfunction

Jayr, C., Garat, C., Meignan, M., Pittet, J.F., Zelter, M., and Matthay, M.A. 1994. Alveolar liquid and protein clearance in anesthetized ventilated rats. J. Appl. Physiol. 76:2636-2642.

Lipke, A.B., Matute-Bello, G., Herrero, R., Kurahashi, K., Wong, V.A., Mongovin, S.M., and Martin, T.R. 2010. Febrile-range hyperthermia augments lipopolysaccharide-induced lung injury by a mechanism of enhanced alveolar epithelial apoptosis. J. Immunol. 184:3801-3813. Matalon, S., Benos, D.J., and Jackson, R.M. 1996. Biophysical and molecular properties of amiloride-inhibitable Na+ channels in alveolar epithelial cells. Am. J. Physiol. 271:L1-L22. Matthay, M.A., Landolt, C.C., and Staub, N.C. 1982. Differential liquid and protein clearance from the alveoli of anesthetized sheep. J. Appl. Physiol. Respir. Environ. Exerc. Physiol. 53:96104. Matthay, M.A., Robriquet, L., and Fang, X. 2005. Alveolar epithelium: Role in lung fluid balance and acute lung injury. Proc. Am. Thorac. Soc. 2:206-213. Mazzon, E. and Cuzzocrea, S. 2007. Role of TNF-α in lung tight junction alteration in mouse model of acute lung inflammation. Respir. Res. 8:75. Modelska, K., Pittet, J.F., Folkesson, H.G., Courtney Broaddus, V., and Matthay, M.A. 1999. Acid-induced lung injury. Protective effect of anti-interleukin-8 pretreatment on alveolar epithelial barrier function in rabbits. Am. J. Respir. Crit. Care Med. 160:1450-1456. Mutlu, G.M. and Sznajder, J.I. 2005. Mechanisms of pulmonary edema clearance. Am. J. Physiol. Lung Cell Mol. Physiol. 289:L685-L695. Pittet, J.F., Brenner, T.J., Modelska, K., and Matthay, M.A. 1996. Alveolar liquid clearance is increased by endogenous catecholamines in hemorrhagic shock in rats. J. Appl. Physiol. 81:830-837. Rezaiguia, S., Garat, C., Delclaux, C., Meignan, M., Fleury, J., Legrand, P., Matthay, M.A., and Jayr, C. 1997. Acute bacterial pneumonia in rats increases alveolar epithelial fluid clearance by a tumor necrosis factor-alpha-dependent mechanism. J. Clin. Invest. 99:325-335. Sakuma, T., Pittet, J.F., Jayr, C., and Matthay, M.A. 1993. Alveolar liquid and protein clearance in the absence of blood flow or ventilation in sheep. J. Appl. Physiol. 74:176-185. Sakuma, T., Folkesson, H.G., Suzuki, S., Okaniwa, G., Fujimura, S., and Matthay, M.A. 1997. Betaadrenergic agonist stimulated alveolar fluid clearance in ex vivo human and rat lungs. Am. J. Respir. Crit. Care Med. 155:506-512. Sartori, C. and Matthay, M.A. 2002. Alveolar epithelial fluid transport in acute lung injury: New insights. Eur. Respir. J. 20:1299-1313. Smedira, N., Gates, L., Hastings, R., Jayr, C., Sakuma, T., Pittet, J.F., and Matthay, M.A. 1991. Alveolar and lung liquid clearance in anesthetized rabbits. J. Appl. Physiol. 70:1827-1835. Suzuki, S., Zuege, D., and Berthiaume, Y. 1995. Sodium-independent modulation of Na(+)K(+)-ATPase activity by beta-adrenergic agonist in alveolar type II cells. Am. J. Physiol. 268:L983-L990.

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Ware, L.B. and Matthay, M.A. 2001. Alveolar fluid clearance is impaired in the majority of patients with acute lung injury and the acute respiratory distress syndrome. Am. J. Respir. Crit. Care Med. 163:1376-1383. Xie, W., Wang, H., Wang, L., Yao, C., Yuan, R., and Wu, Q. 2013. Resolvin D1 reduces deterioration of tight junction proteins by upregulating HO-1 in LPS-induced mice. Lab. Invest. 93:991-1000. Yue, G. and Matalon, S. 1997. Mechanisms and sequelae of increased alveolar fluid clearance in hyperoxic rats. Am. J. Physiol. 272:L407-L412.

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How to measure alterations in alveolar barrier function as a marker of lung injury.

The alveolar capillary membrane maintains the proper water and solute content of the epithelial lining fluid at the alveolar air-liquid interface, whi...
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