An Optimized Evans Blue Protocol to Assess Vascular Leak in the Mouse

Summary

In this article, an economical, optimized, and simple protocol is described which uses the Evans blue dye method for assessing plasma extravasation in the organs of FVBN mice that can be adapted for use in other strains, species, and other organs or tissues.

Abstract

Vascular leak, or plasma extravasation, has a number of causes, and may be a serious consequence or symptom of an inflammatory response. This study may ultimately lead to new knowledge concerning the causes of or new ways to inhibit or treat plasma extravasation. It is important that researchers have the proper tools, including the best methods available, for studying plasma extravasation. In this article, we describe a protocol, using the Evans blue dye method, for assessing plasma extravasation in the organs of FVBN mice. This protocol is intentionally simple, to as great a degree as possible, but provides high quality data. Evans blue dye has been chosen primarily because it is easy for the average laboratory to use. We have used this protocol to provide evidence and support for the hypothesis that the enzyme neprilysin may protect the vasculature against plasma extravasation. However, this protocol may be experimentally used and easily adapted for use in other strains of mice or in other species, in many different organs or tissues, for studies which may involve other factors that are important in understanding, preventing, or treating plasma extravasation. This protocol has been extensively optimized and modified from existing protocols, and combines reliability, ease of use, economy, and general availability of materials and equipment, making this protocol superior for the average laboratory to use in quantifying plasma extravasation from organs.

Introduction

Vascular leak in the organs refers to extravasation, or leakage of blood plasma through gaps produced in the endothelium of post capillary venules in the organs. This plasma extravasation or increased vascular permeability, which may arise from some type of an inflammatory response, may have grave consequences. Thus, it is important that this phenomenon, its causes, modulators, and consequences, are studied and understood, and likewise, that investigators have good tools and protocols with which to study them. The endothelial gaps may be produced via a number of stimuli, but usually are produced by the action of peptide neurotransmitters and/or tachykinins on the endothelia. One of the major naturally occurring mediators of this process, which results in increased plasma extravasation, is the undecapeptide tachykinin neuropeptide, substance P¹.

Methods to investigate and measure vascular permeability or plasma extravasation, which use the albumin-binding property of Evans blue dye, have been developed, and are usually known for their accuracy, simplicity, economy, safety, and ability to allow the determination of plasma extravasation from several tissues at once, if so desired^{2,3,4,5,6,7,8,9}. This Evans blue protocol for assessing plasma extravasation in the organs of FVBN mice uses all these, but adds some important modifications that make it generally useful and adaptable for future studies, involving the average laboratory that conducts or will conduct important studies of factors associated with plasma extravasation or vascular permeability. In this protocol, substance P is introduced to the mice at 1 nmol/kg, which augments the extravasation of plasma by 1.5-fold. This increases the sensitivity of the protocol, resulting in more easily observable and obtainable results. Other factors that impact permeability, such as various other peptides, chemicals, or some forms of toxic injury, may be used or studied by other laboratories, as desired. Jugular vein injections are used in this protocol to introduce Evans blue and substance P systemically, which requires terminal surgery. However, jugular vein injections^5,7,10, even after consideration of the necessary terminal surgical techniques, are easier to master and lead to the production of more consistent results than other venous injections, including tail vein injections^4,9. Although it may be possible for Evans blue to be delivered by retro-orbital venous sinus injections, no references in the literature have been found that use this method of delivery of Evans blue. However, as for tail vein injections, the high degree of expertise and practice to reproducibly master this technique greatly limits its use for successful Evans blue injections. In contrast, the alternative jugular vein injection method as described in our protocol, offers a technically obtainable solution. A crucial procedure for perfusion of the mouse's veins, performed just after the sacrifice of the Evans blue-perfused mouse, removes excess Evans blue dye, and has been standardized in this protocol. Previously described methods of perfusion have been carefully examined and modified to obtain the present procedure.Other modifications described here are all optimized, straightforward, and inexpensive.

There are some important limitations of the Evans blue dye method. For example, low sensitivity sometimes associated with this method may prevent some additional gross pathological and histological examination of tissues from Evans blue-injected animals. However, these and other limitations have led to the development of alternative methods and models that, nonetheless, still use Evans blue. The measurement of Evans blue by fluorescence (rather than by visual-range) spectroscopy may increase the sensitivity of the method. Additionally, fluorescence microscopy of Evans blue-stained tissues was developed to allow for observation of vascular leak in more distinct locations¹¹. Also, whole-body imaging and scanning of a live animal previously injected with Evans blue¹² allows for investigation of Evans blue concentrations in a continuous manner, rather than at one specific chosen time point of the experiment. However, this method requires the availability of appropriate imaging facilities, and may be very expensive. Modifications involving Evans blue and performed in an in vitro type of model, such as in a cell culture or chick chorioallantoic model¹³ (CAM) have also been described. These models are monitored by fluorescence and intravital¹⁴ microscopy, and allow the quantification of vascular permeability changes over time, but may raise questions regarding accurate modeling of in vivo conditions and may also be expensive.

There have been other methods developed to determine and quantify vascular leak or permeability, which do not involve the administration of Evans blue. These methods may employ an appropriate fluorescent molecule (such as albumin or fluorescein), or an isotopically labeled or otherwise tagged molecule, to live animals (or to cell culture or chorioallantoic (CAM) models¹³, followed by non-invasive imaging (PET scanning, MRI, intravital microscopy, whole body scanning) or by invasive imaging (fluorescent microscopy)^3,12,15. Although these techniques may offer a number of advantages over other Evans blue methods, they also have disadvantages, which may include their considerable complexities, requisite expertise, resources, and high monetary costs.

Neprilysin¹⁶ (the peptidase enzyme NEP, also known as CD10, MME, or Enkephalinase) has been suggested to be involved in inhibiting plasma extravasation, at least in part, through the enzymatic metabolism and inactivation of endogenous substance P. Thus, in tissues in which the cell surface peptidase NEP occurs, there may be an attenuation of the effect of substance P, presumably by the peptidase activity of NEP.

Initially, we tested for substance P-induced plasma extravasation utilizing this modified Evans blue protocol, with FVBN wild type (WT) and NEP knockout (KO) mice. NEP involvement in substance P-augmented plasma extravasation was suspected from these initial studies, and we describe these and further experiments involving NEP's role in plasma extravasation. However, the focus of this manuscript is not NEP or its role in plasma extravasation, but rather the plasma extravasation experiments themselves. The NEP results are representative of the kind of results that may be obtained through use of this modified protocol. The Evans blue method to measure plasma extravasation has been optimized and modified, as described in detail below for FVBN mice.

Protocol

All applicable international, national, and/or institutional guidelines for the care and use of animals (mice) were followed in the experiments described in this manuscript.

This method uses FVBN adult mice, aged 16 - 20 weeks, found to be optimal for the purposes of this study. Day 1 includes steps 1 - 5 and Day 2 includes steps 6 - 7 (Figure 1).

1. Equipment Preparation

1. Secure an adequate supply of sterile, disposable syringes and needles, if ketamine/xylazine is used as the anesthetic (as recommended). If isoflurane is used as the anesthetic, check the oxygen tank and the fluid level of isoflurane to make sure there are adequate supplies for the experiment before starting. Also, assemble the nosecone breathing circuits and attach them to the induction box; attach new charcoal canisters to the breathing circuits. Prepare the induction box by turning on the oxygen and ascertaining that the second stage reads approximately 50 psi.
2. Turn on the heating pad to 37 °C.
3. Prepare the rectal temperature probe for the surgery.

2. Mouse Preparation

This step includes anesthesia, hair removal, and positioning (adult FVBN mice-age 16 - 20 weeks).

1. Weigh the mice and record the weights.
2. Anesthetize the mice.
   1. Administer ketamine and xylazine IP (80 - 100 mg/kg and 7.5 - 16 mg/kg, respectively). However, it is recommended to begin with lower doses of ketamine and xylazine (about 30 mg/kg and 6 mg/kg, respectively).
   2. Maintain the anesthesia with about 0.1 - 0.25 times initial doses of ketamine/xylazine throughout the surgery. Ketamine and xylazine were the anesthetic agent(s) of choice in this particular study, as more reproducibility and survivability were observed. The anesthetic agent(s) of choice in other studies may be found to be different.  If isoflurane is used, put the mouse in an induction chamber and turn on isoflurane to 5% until the mouse loses full consciousness. Then use a nasal nosecone set at 1.5 - 2.5% isoflurane throughout the remainder of the procedure.
3. Monitor the mice every 2 - 3 min by toe pinch to check for appropriate depth of anesthesia.
4. Shave the ventral neck area of the mouse.
5. Place the mouse in a supine position on the preheated pad. Secure the paws and feet of the mouse to the surgical surface with tape.
6. Place artificial tear ointment onto the eyes to prevent drying out during surgery.

3. Surgical Details

1. Make a 1 cm incision in the right ventral neck over the jugular vein.
2. Apply one or two drops of lidocaine (1 - 2%) into the incision area for pain management and to promote vasodilation. Wait 2 min. for the lidocaine to take effect. 
3. Expose and isolate the right internal jugular vein via blunt dissection. Tie the vein off with 4-0 suture and gently retract the rostral end of the vessel with a hemostat. Cut a hole, using fine scissors, about 3 mm below or caudal to the tie, approximately halfway through the diameter of the vein.
4. Mark a PV-1 polyvinyl catheter 1.5 cm from the end and insert it into the jugular vein through the hole using a vessel dilator, and thread the catheter towards the caudal end of the vessel, approximately 1.5 cm.
5. Tie the catheter securely within the caudal part of the vessel (below the cut), with 4-0 silk. Tie the rostral part of the vessel to the outside of the catheter with the loose ends of the suture which was used to tie off the rostral jugular vein, in step 3.3.
6. Tack the skin loosely back together around the catheter with 4-0 silk to help prevent loss of body heat and desiccation of tissues.
7. Connect the catheter to a syringe containing heparinized saline (10 U/mL) and flush the catheter.

4. Injections

1. Inject the Evans blue solution (50 μL of a 30 mg/mL solution in 0.9% normal, unheparinized saline, or approximately 50 mg/kg) into the jugular vein catheter, followed by a small volume of heparinized saline to flush the line.
2. 2 min later, inject substance P (100 μL of a 0.3 μM solution in 0.9% normal, unheparinized saline, or 1 nmol/kg), followed by a small volume of heparinized saline to flush the line. Substance P augments the extravasation of plasma proteins through the endothelial layer; in this protocol, it routinely induces an augmentation of plasma extravasation values of approximately 1.5-fold, making plasma extravasation values easier to measure.
3. Wait 18 min after the substance P is injected. During this time, the Evans blue dye will equilibrate and circulate.
4. Terminate the experiment 18 min after the injection of substance P (20 min after Evans blue injection) by sacrificing the mouse with cervical dislocation. It is likely that the mouse can be directly cervically dislocated, without first giving an overdose of anesthetic, as the mouse is probably still well-anesthetized from the surgery. However, if it is necessary, an overdose of the anesthetics ketamine/xylazine, isoflurane, or pentobarbital may be given, followed by cervical dislocation.

5. Isolation of Organs

1. Cut open the chest cavity of the mouse and gravity-perfuse (from a height of about 51 cm or 20") the heart and blood vessels with 50 mL of 50 mM sodium citrate, pH 3.5. pH 3.5 presumably preserves Evans blue binding to albumin.
2. Excise 1 - 5 relevant organs (tissues) with a dissecting scalpel (e.g. urinary bladder, kidney, stomach, liver, pancreas, proximal or distal colon, ileum, duodenum, flank skin, ears, tail, heart, and/or lungs) and remove any residual contents, if present.
3. Rinse the organs in room temperature (RT) phosphate-buffered saline (PBS; 1.44 g of Na₂HPO4, 0.24 g of KH₂PO4, 8.0 g of NaCl, and 0.20 g of KCl in 1 L, pH 7.4).
4. Blot the organs with tissue, cut each organ in half, and weigh each half (wet weights, in g).
5. Dry one half of the tissue in a drying oven at 150 °C, on foil, for 48 h.
6. Place the other tissue half in a consistent volume (up to 200 µL) of formamide in a microfuge tube for 48 h (and up to 72 h) to extract the Evans blue.

6. Measurement of Tissue OD

1. Remove 50 µL of Evans blue-infused formamide (after 48 - 72 h RT incubation) from the microfuge tube and place into one well of a 96-well polystyrene plate. Be careful not to transfer tissue pieces along with the formamide.
2. Place 50 µL of new, pure formamide into each of two empty wells of the 96-well plate for the blanks.
3. Measure and record the OD₆₂₀ of each well of the 96-well plate on an absorbance plate reader. 620 nm is the absorbance max of Evans blue.

7. Calculation of Plasma Extravasation

1. Weigh the dry tissue half which has been in the oven for 48 h.
2. Calculate the wet weight/dry weight ratio for the specific organ of interest from each individual mouse, starting with the wet weight of this tissue half (obtained in step 5.4), divided by the dry weight of this same tissue half (obtained in step 7.1).
3. Calculate the dry weight (in g) of the tissue half in formamide by dividing the wet weight of the tissue half before it was placed in formamide (obtained in step 5.4) by the wet: dry weight ratio for the specific organ of interest (calculated in step 7.2).
4. Calculate the corrected OD₆₂₀ values. Starting with the OD₆₂₀ value from each experimental well of the 96-well plate (containing Evans blue-infused formamide from each tissue, obtained in step 6.3), subtract the blank well OD₆₂₀ value (the mean OD₆₂₀ value of the two wells containing pure formamide, prepared in step 6.2, OD₆₂₀ values obtained in step 6.3) from each experimental value.
5. Calculate the plasma extravasation value by dividing the corrected OD₆₂₀ (calculated in step 7.4) by the dry weight of the tissue half in formamide (calculated in step 7.3). The units of plasma extravasation will be OD₆₂₀/g dry weight.
6. Analyze data and express as the mean ± SEM. Statistically compare groups by t test or one-way analysis of variance and Scheffé's multiple-comparison test (lycofs01.lycoming.edu), as appropriate.

Results

In Figure 1, a schematic of the procedure is shown, which has been found to result in the most reliable and consistent substance P-induced plasma extravasation values from organs of FVBN mice. This procedure usually takes two days of work, separated by at least 48 h of waiting time. It is possible to spread it out even more, if this is done consistently for all experiments to be compared. For example, after the organs are isolated on Day 1, the organs can be ...

Discussion

As discussed above, the study of plasma extravasation may ultimately lead to new knowledge concerning the causes of or new ways to inhibit or treat plasma extravasation. The successful use of the plasma extravasation protocol (above), using Evans blue dye, has been demonstrated in the current manuscript. Although the data shown back the hypothesis that NEP may protect the vasculature against plasma extravasation, this is a secondary goal presently, with the primary goal being to present an optimized protocol which may be...

Materials

Name	Company	Catalog Number	Comments
isoflurane	Vet One	200-070	inhaled anesthetic
ketamine	Vet One	200-055	injectable anesthetic
xylazine	Lloyd Laboratories	139-236	injectable anesthetic
syringes (10,3 & 1 cc)	Becton Dickinson	309604, 309657, 309659
needles (20G1,23G1 & 26G1/2)	Becton Dickinson	305178, 305193, 305111
isoflurane induction chamber	VetEquip	941443	1 Liter
nosecone breathing circuits	VetEquip	RC2	Rodent Circuit Controller 2
oxygen tank	Airgas	UN 1072	100% medical
heating pad	CWE Inc.	TC-1000	temperature controller
rectal temperature probe	CWE Inc.	10-09012	mouse
balance (for rodents)	Ohaus	CS 2000
surgical tools-scissors	Fine Science tools	15000-00	Vannas Spring scissors 3mm straight blade (cutting vessels)
surgical tools-forceps	Fine Science tools	11151-10	Graefe extra fine forceps (isolating mouse vessels)
surgical tools-hemostats	Fine Science tools	13009-12	Halstead-mosquito hemostats (blunt dissect, hold tissue)
surgical tools -suture drivers	Fine Science tools	12502-12	Olsen-Hegar suture drivers (suturing)
surgical tools-forceps	Fine Science tools	11627-12	Adson-Brown alligator forceps (tissue grasping suturing, rat)
surgical tools-scissors	Fine Science tools	14110-15	Mayo tough cut scissors 15 cm (surgery, dissection, bones, rat)
surgical tools-forceps	Fine Science tools	18025-10	suture tying forceps (used for Millar cath)
surgical tools-scissors	Fine Science tools	14078-10	Lexer Baby scissors straight (surgery, mouse)
surgical tools-forceps	Fine Science tools	11254-20	Dumont #5 fine-tip forceps (rat vessels, dissection)
surgical tools-scissors	Fine Science tools	14082-09	Dissector scissors 12 mm (surgery, rat mouse)
surgical tools-forceps	Fine Science tools	11051-10	10 cm Graefe forceps (tissue grasping, rat mouse)
surgical tools-forceps	Fine Science tools	11251-35	Dumont 5/45 forceps (introducer for vessels)
surgical tools-retractors	Fine Science tools	17012-11	Weitlaner retractors 2/3 tooth (rat surgical)
surgical tools-forceps	Fine Science tools	11294-00	Dumont #4 forceps (vessel isolation rats, mice)
surgical tools-forceps	Fine Science tools	11297-00	Dumont #7 forceps (tissue grasping, dissection)
surgical tools-scissors	Fine Science tools	14058-11	tough cut iris scissors (mouse dissection, bones)
surgical tools-forceps	Fine Science tools	11009-13	serrated, curved Semken forceps (tissue grasping, mouse rat)
surgical tools-hemostats	Fine Science tools	13003-10	Hartman curved hemostats (blunt dissect, hold tissue)
surgical tools-forceps	Fine Science tools	11006-12	Adson serrated forceps (tissue grasping)
clippers	Oster	A5
tape	Fisherbrand	159015G
artificial tear ointment	Akorn Inc	13985-600-03
lidocaine	Hospira	0409-4277-01	2% injectable
polyvinyl catheters	Tygon	PV-1
Evans blue	Sigma Aldrich	E2129
Substance P	Bachem	H-1890
heparin	Sagent Pharmaceuticals	25201-400-10	1000 U/ml
saline solution	Hospira	0409-7138-09	0.9% sodium chloride
phenobarbital	Vortech	0298-9373-68
sodium citrate	Fisher Scientific	BP327-1
PBS	Sigma Aldrich	P4417-50TAB
Kimwipes for blotting	Fisher Scientific	06-666A
formamide	Sigma Aldrich	47670
microbalance	Denver Instrument	APX-60
microfuge tubes	Fisher Scientific	07-200-534
polystyrene 96 well plate	Becton Dickenson	351172
absorbance plate reader	BioTek	Synergy 2
polyacrylamide gels	Bio-Rad	3450014
protein molecular weight standard	Bio-Rad	1610374
Protran supported nitrocellulose	Amersham (GE)	10600015
gel box	Bio-Rad	1658005
Tris	Fisher Scientific	BP152-1
Tween20	Sigma Aldrich	P-1379
sodium chloride	Fisher Scientific	S271-1
primary NEP polyclonal antibody	R & D Systems	AF1182
doxycycline chow	Teklad (HARLAN)	TD.130750
FVB/NJ wild type mice	Jackson	001800
secondary antibody (goat anti-rabbit)	ZyMed	81-6120
ECL solution-Western Lightening Plus	PerkinElmer	NEL104001EA
film	Pierce	34091