Measurement of lung vessel and epithelial permeability in vivo with Evans Blue

Abstract

Lung fluid balance is maintained in part by the barriers formed by the pulmonary microvasculature and alveolar epithelium. Failure of either of these barriers leads to pulmonary edema, which limits lung function and exacerbates the severity of acute lung injury. Here we describe a method using Evans Blue dye to simultaneously measure the function of vascular and epithelial barriers of murine lungs in vivo.

1. Introduction

In order to efficiently mediate gas exchange, the lung needs to efficiently control fluid balance. Injury and inflammation impair the control of lung fluid, resulting in pulmonary edema which, when extensive, exacerbates lung injury and causes acute respiratory distress syndrome (ARDS) [1]. Control of the lung air/liquid barrier is predominantly mediated by two distinct systems, the pulmonary microcirculation [2,3] and the alveolar epithelium [4], failure of which can lead to interstitial fluid accumulation (tissue edema) and airspace flooding, respectively (Figure 1). Tissue edema and airspace flooding both have the capacity to impair gas exchange and exacerbate injury. Understanding the relative contributions of the vascular endothelial and alveolar epithelial barriers in regulating lung fluid is critical to identifying effective therapeutic approaches to the treatment of ARDS.

Figure 1. Diagram depicting a cross-section though a normal lung alveolus comprised of an epithelial sac surrounded by blood vessels.

Note that the transit of fluid-borne molecules from the vasculature must traverse across the vascular endothelial barrier though the interstitial tissue and across the alveolar epithelial barrier. Reproduced from [12] with permission.

It has long been appreciated that Evans Blue dye (T-1824) preferentially binds to serum albumin [5], making it an effective marker for the ability of albumin to extravasate across barriers and accumulate in tissues. Albumin is a 68 kDa, native serum protein that has been found to penetrate barriers and accumulate in airspaces in response to lung injury (e.g., [6]), which underscores its utility as a marker for lung barrier failure. Albumin permeability also is a particularly useful marker in that it can cross endothelial and epithelial barriers by both the paracellular route through tight junctions [7] and the transcellular route through transcytosis [8–10].

In contrast to direct measurement of albumin in different lung compartments by ELISA or immunoblot, Evans Blue is easily measured by absorbance spectroscopy and is highly sensitive [11]. This is particularly critical for measurement of dye accumulation in the interstitium. Here we describe a protocol that enables the accumulation of Evans Blue in the airspace and interstitial compartments to be simultaneously determined, in order to measure the relative impact of injury on lung endothelial and epithelial barriers.

2. Materials

2.1. Evans Blue Injection

1. Dulbecco’s Phosphate Buffered Saline (DPBS) without Ca²⁺/Mg²⁺ (Corning/Mediatech #21–031-CV, Manassas, VA)

2. Evans Blue: Make a 0.5% solution in DPBS by adding 0.05 g of Evans Blue (MilliporeSigma E2129, St. Louis, MO) to 10 ml of DPBS (w/o Ca²⁺/Mg²⁺), then filter sterilize. Store at room temperature protected from light.

3. Sedative/Anesthetic cocktail (20% Ketamine + 5% xylazine solution): Add 1 ml 100 mg/ml ketamine (Mylan Institutional NDC 67457-108-10, Galway, Ireland). and 0.25 ml 20 mg/ml xylazine (Zoetis Inc NDC 59399-110-20, Kalamazoo, MI) to 3.75 ml DPBS without Ca²⁺/Mg²⁺ to produce 5 ml total solution

4. Isoflurane (Piramal Enterprises Limited NDC 66794-017-25, Telangana, India)

5. 25G × 5/8 needles (BD PrecisionGlide #305122, Franklin Lakes, NJ) and 30G × 1 needles (BD PrecisionGlide #305128) (see Note 1).

6. 1 ml syringes (BD #309659)

7. Cotton Gauze Sponges, 5 × 5 cm (FisherBrand #22-362-178, Pittsburgh, PA)

8. Heat lamp

9. Small animal heating pad

2.2. Tissue Harvest

1. Animal surgical instruments: dissecting scissors, fine tip tweezers, smooth and rat tooth forceps, dissection board.

2. Portable balance to weigh animals (Mettler Toledo PL6000-S or equivalent)

3. Mouse trachea tube (DWK Kimble Kontes Brand Microflex Syringe Needles-Blunt, Fisher Scientific K868280–2001)

4. 5–0 Silk Suture (Ethicon #K870H,Somerville, NJ).

5. 21G × 1 ½ needles (BD PrecisionGlide #305167).

6. 10 ml syringe (BD #302995).

7. 1.5 ml conical microfuge tubes (Eppendorf, # 022364111, Enfield, CT). Each mouse requires 3 collection tubes each for blood/serum, bronchoalveolar lavage (BAL) fluid, and lung tissue.

8. Heparin (Aurobindo Pharma Limited NDC 63739-920-25, Memphis, TN) stock concentration 1000 USP: Used to coat needle and syringe for blood collection.

9. Formamide (Electron Microscopy Sciences #15745, Hatfield, PA). Add 250 μl per tube for extraction of lung tissue

10. DPBS with Ca²⁺/Mg²⁺ (Corning/Mediatech #21–030-CV)

11. Lavage solution: DPBS with Ca²⁺/Mg²⁺ containing 1:200 200mM phenylmethylsulfonyl fluoride (PMSF, MilliporeSigma, # P7626) and 1:500 1 M NaF (MilliporeSigma #S7920): Prepare 500 μl per mouse and keep lavage fluid on ice in an ice bucket (see Note 2)

2.3. Tissue Analysis

1. Heating block (Fisher Scientific Dry Bath Incubator #11–718 or equivalent)

2. Refrigerated microfuge (Eppendorf 5415 R or equivalent)

3. Microplate reader (Biotek Synergy H1 Multimode Plate Reader or equivalent)

3. Methods

3.1. Tail Vein Injection of Evans Blue

1. Prepare 0.5% Evans Blue solution.

2. Place mice in a cage under the heat lamp. Warm mice for 5 −10 min in order to dilate blood vessels (see Note 3).

3. Weigh each mouse and intraperitoneally (IP) inject 10 μl/g body weight + 10 μl ketamine/xylazine anesthetic cocktail. (see Note 4).

4. Place the mice back in the cage and allow anesthetic to take effect (see Note 5).

5. Place mouse on a platform so that the mouse is at a comfortable height and the tail hangs over the edge.

6. Rotate the tail to locate the lateral tail vein and potential injection site. Make sure the vein is facing upward (Figure 2).

7. Clean injection area with warm alcohol pad.

8. Fill syringe with 400 μl Evans Blue solution making sure to avoid air bubbles (see Note 6).

9. Hold the tail with the nondominant hand while stretching the tail so that the injection site is visible and the vein is in a straight line.

10. Insert 30G needle at a 10–15° angle, bevel up, into the injection site. Advance needle towards the head keeping the needle and syringe parallel to the tail. The needle should be visibly in the tail vein (Figure 2).

11. Slowly inject 200 μl of dye into the tail vein of the mouse. If you have correct placement of the needle the plunger will advance with ease (see Note 6).

12. Place mouse on heating pad and wait 1 h (see Note 7). Evans Blue will clearly dye the nose and footpads of treated mice (Figure 3).

Figure 2. Tail vein injection.

(a,b) Diagram showing mouse tail vasculature. Branch vessels extend from the artery to the veins. Red arrows, artery; blue arrows, vein; green arrows, branch vessels. (c-f) Procedure for tail vein injection. (c) The outer tube is grasped by the first and second fingers. The third finger is placed under the inner cylinder. Place the needle on the surface of the tail in parallel (d) and insert it carefully (e). Once the needle tip is under the skin, pull back the syringe slightly during insertion to confirm that blood flows back to ensure that a vein is penetrated (f, arrow). Reproduced from [13] with permission.

Figure 3. Appearance of Evans Blue in mice and murine lungs.

a,b Mice that were either untreated (a) or intravenously injected with Evans Blue (b). Sixty minutes after injection, the treated mice show clear blue labeling of the nose and paws. (c-e). Representative images of lung tissue from mice that were either untreated (c), or after Evans Blue tail vein injection showing control (d) or alcohol fed and endotoxin treatment to induce acute lung injury (e). Uninjured mice show little tissue accumulation (arrow) of Evans Blue as compared with injured mice.

3.2. Serum Collection

1. Rinse needle (21G) and 1 ml syringe with heparin in order to coat both needle and syringe (see Note 8)

2. Sacrifice the mouse using an approved injectable anesthetic overdose procedure. A 1 ml IP injection 1 ml of 20% Ketamine + 5% xylazine solution is sufficient (see Note 9).

3. Once unresponsive, pin mouse legs down on dissection board. Immobilize the head by placing a suture loop around the front incisors and pinning the loop down to the dissection board (see Note 10).

4. Wet the abdomen with water or ethanol to prevent fur from entering the abdominal cavity and contaminating tissue. Cut open the skin and peritoneum along the midline.

5. Cut though the ribcage and open chest cavity so that the bottom of the heart is exposed.

6. Puncture the left ventricle with the 21G needle attached to the heparinized 1 ml syringe. Slowly draw plunger back into syringe to start collecting blood. Collect a minimum of 250 μl blood/mouse.

7. Evacuate blood into collection tube and place on ice. (see Note 11)

8. Centrifuge blood 3,000 × g at 4°C for 15 min. Collect supernatants (serum) and transfer to a new tube. Freeze at −20°C (see Note 12).

3.3. BAL Fluid Collection

1. Cut away skin on the neck so that the area near the trachea is exposed. Carefully snip away thin tissue covering the trachea (see Note 13).

2. Turn the mouse so that the head is closer to you.

3. With the help of forceps, lay a string of surgical sutures behind the trachea.

4. Make a small nick at a 45° angle on the top surface of the trachea nearest the larynx (see Note 14).

5. Slide the 20G trachea tube into the nick and stop right above the ribcage. Ensure the surgical string is at least 2–3mm above the bottom tip of the trachea tube.

6. Secure the needle in place by tying a knot with the sutures around it.

7. Remove any air in the lungs by evacuating at least 1 ml of air from the lung using an empty 1 ml syringe.

8. Quickly replace the empty syringe with one filled with 500 μl of lavage fluid and slowly fill lungs with the fluid. Rinse the lungs twice with the same lavage fluid, then pull the plunger out to retrieve the final volume (see Note 15).

9. Collect the fluid in centrifuge tubes and place on ice.

10. Centrifuge BAL at 3,000 × g at 4°C for 15 min. Collect supernatants and transfer to a new tube. Freeze at −20°C (see Note 12).

3.4. Lung Tissue Collection

1. Rotate the mouse back to the original position with the feet closest to you.

2. Snip the aorta below the diaphragm so that blood can easily move from the perfused lungs (see Note 16).

3. Take a DPBS with Ca²⁺/Mg²⁺-filled 10 ml syringe attached to a 25G needle and insert the needle into the right ventricle, aiming in the direction of the pulmonary artery. Gently push the plunger of the syringe to perfuse the lungs with 5 ml of DPBS (see Note 17).

4. Carefully dissect out the lungs from the thoracic cavity by grabbing the trachea with forceps then snipping down behind the lungs while gently pulling the lungs away.

5. Once out, rinse the exterior of the lungs with DPBS with Ca²⁺/Mg² to remove any clotted exterior blood and cut away extra-pulmonary tissue (e.g., heart and vasculature) and remove cartilaginous tissue (trachea and large bronchi). Lungs with tissue accumulation of Evans Blue are visibly stained (Figure 3c–e).

6. Place half the isolated lung tissue in a centrifuge tube containing 250 μl of formamide on ice. Weigh lung samples and record for later analysis (see Note 18).

7. Incubate at 55°C in a heating block for 48 h.

8. Centrifuge lung tissue at 3,000 × g at 4°C for 15 min. Collect supernatants and transfer to a new tube.

3.4. Evans Blue Measurement and Analysis

1. Aliquot 100 μl/well of samples from serum (diluted 1:10 in PBS), undiluted BAL Fluid, and lung tissue extract in a 96-well plate (see Note 19).

2. Measure absorbance of samples and standards at 620nm and 740nm using a microplate reader (see Note 20).

3. Correct the A₆₂₀ readings for turbidity in BAL fluid and lung tissue using the correction factor y = 1.193x + 0.007 where x is A₇₄₀ and A₆₂₀ corrected = yA₆₂₀ [11]. Use standards to get absolute Evans Blue concentrations in μg/ml.

4. Divide BAL fluid Evans Blue concentration by serum Evans Blue concentration to normalize and account for variability in the tail vein injections (see Note 21).

5. Divide lung tissue Evans Blue concentration by lung weights and then by serum Evans Blue concentration (see Note 22).

4. Notes

1. 30G needles are used for tail vein injection but 25G needles are used for sedation, since 30 G needles are too small to quickly sedate mice at the volumes required.

2. Pre-cooling lavage fluid on ice before BAL extraction significantly increases the yield and helps inhibit protease activity when harvesting BAL.

3. Make sure that the heat lamp is not too close to the cage by placing your hand near the bottom of the cage and holding for 30 s. The lamp should be no closer than 20–30 cm from the mice. If the heat is uncomfortable to you the lamp is too close and may cause the mice to overheat.

4. This anesthetic guideline is for mice weighing 25 – 35 g. For mice smaller than 25 g, use 10 μl/g body weight ketamine/xylazine. For mice larger than 35 g, use 10 μl/g body weight + 15 μl ketamine/xylazine. Alternatively, Evans Blue injection can be done on mice without anesthetic using a mouse restraint device (Red Tailveiner Restrainer Braintree Scientific Inc. #TV-RED 150-STD, Braintree, MA). Note that sedated and nonsedated mice will give different Evans Blue permeability values due to differences in heart rate, breathing rate and stimulation of the sympathetic system.

5. Mice usually are non-responsive to a foot pinch after 5 min as an indication that they are anesthetized.

6. Although the syringe is filled with 400 μl Evans Blue solution, only 200 μl is administered. Having an excess of dye solution in the syringe enables the amount injected to be more accurately monitored, especially if the tail vein is initially missed. If it is difficult to advance the plunger the needle is not in the tail vein. The entire mouse should turn blue within 1 min, most notably in the pads of the feet, the nose, and the ears

7. The heating pad step is only required if anesthetic was used. The heat keeps mice warm as they recover to avoid a drop in body temperature due to the anesthetic. If mice were not anesthetized, place the mouse back into the cage for 1 h.

8. After 1 h mice are ready for tissue harvest. Harvest in the following order: 1) blood, 2) BAL fluid, 3) lung tissue.

9. Overdose with ketamine/xylazine is the preferred method of euthanization since CO₂ asphyxiation stops the heart from beating and reduces recovery of blood during collection. Harvest blood immediately after sacrifice for maximum collection. Do not use cervical dislocation, since this can tear the trachea rendering mice unable for BAL collection.

10. The head should be tilted back fully with the neck fully exposed.

11. It can be difficult to distinguish serum from red blood cells because they will be heavily dyed with Evans Blue. By collecting at least 200 μl of whole blood, the top 60 μl supernatant can be isolated without disrupting the cellular layer on the bottom.

12. Serum and centrifuged BAL fluid are stored frozen prior to analysis since the lung tissue requires 48 h processing time.

13. The trachea is beneath the salivary glands. Glands can be carefully cut away and gently pulled apart to uncover the trachea.

14. The nick should only cut the top surface of the trachea and should be just large enough to accommodate the 20G trachea tube. The cut should open less than half of the diameter of the trachea.

15. The lungs should clearly inflate with the addition of fluid. Typical fluid recovery is 300–350 μl of lavage fluid. This provides enough material for duplicate absorbance measurements and protein determination by the Bicinchoninic Acid (BCA) assay (Sigma-Aldrich #BCA1–1KT).

16. Snipping the aorta reduces the pressure needed when perfusing with PBS into the right ventricle, avoiding potential damage to the lung microvasculature.

17. Lungs should turn white (with patches of blue dye) in color with good perfusion. It is imperative that the lung is properly perfused so that dye from the blood will not interfere with the tissue analysis (Figure 3c–e).

18. Before adding lung tissue, weigh the tubes containing formamide solution alone. This enables net lung weight to be calculated and used for data analysis. Be consistent in analyzing the right or the left lung tissue/lobes. The remaining lung tissue can be snap frozen in liquid nitrogen and used for other analysis (e.g., immunoblot, Q-PCR).

19. Serum must be diluted 1:10 before analyzing to avoid crosstalk with other serum components. BAL and lung tissue extract should not be diluted. When possible, measure absorbance from duplicate or triplicate technical replicates.

20. Standards are made from serial dilutions of Evans Blue stock into DPBS with Ca²⁺/Mg²⁺ (for serum and BAL fluid) or formamide (for lung tissue samples).

21. In this example (Figure 4), mice were given either a control or alcohol diet for 8 weeks and then challenged by either IT or IP lipopolysaccharide (LPS; endotoxin) as a sterile inflammatory challenge [12]. Alcohol rendered the airspaces prone to flooding even in the absence of an additional insult. Interestingly, LPS decreased Evans Blue permeability into the airspaces, most likely due to an increase in Transforming Growth Factor (TGF)-alpha dependent stimulation of Epidermal Growth Factor (EGF) receptors [14]. IT injury, but not IP injury, of alcohol-fed mice also resulted in failure of the endothelial barrier as indicated by the accumulation of Evans Blue in the interstitial space, consistent with increased severity of direct vs. indirect lung injury on lung fluid balance.

22. Assessing barrier function using Evans Blue dye tracks albumin, which is a 68 kDa protein. While this approach offers many advantages, it represents one parameter when considering the effects of injury on lung fluid balance. For instance, macromolecule permeability may not be impacted under conditions where fluid balance is altered by the effects of injury on ion homeostasis [15].

Figure 4. Effect of alcohol consumption and lipopolysaccharide on Evans Blue accumulation in lung airspaces and tissue.

Mice received an IP (a,c) or IT (b,d) treatment with either vehicle control or 5 mg/kg LPS in PBS. After 24 h the mice were administered Evans Blue by tail vein injection and allowed to recover for 2 h. Evans Blue dye was first collected by bronchoalveolar lavage (BAL) (a,b). Lung tissue was then harvested after BAL fluid collection and the right ventricle of the heart was perfused with PBS. Evans Blue dye was extracted from lung tissue by incubating in formamide at 55 °C for 48 h. Evans Blue concentration was analyzed via spectrophotometry (620 nm). Results were corrected for the presence of heme and normalized to Evans Blue serum levels and lung tissue weight. (a,b) Alcohol-feeding alone increased Evans blue levels in BAL fluid, suggesting that alcohol promotes alveolar epithelial barrier dysfunction (n = 6 – 9, ****p < 0.0001; **p = 0.0021). (c,d) Tissue Evans Blue dye content was only significantly increased in alcohol-fed mice that were given an IT administration of LPS. There was no significant change in any other groups. n = 4 – 8, * p = 0.019. Values reported as mean ± standard deviation. Reproduced from [12] with permission.