The authors wish to thank Andy Poczobutt and Dr. Jori Leszczynski for their valuable help and edits to this manuscript.&#160;Supported by Grants received from the National Heart, Lung and Blood Institute (NHLBI RO1 HL078929, PPG HL014985 and RO3 HL095439) and the Department of Veterans&#39; Affairs (Merit Review).

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
<ol>
	<li>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.</li>
	<li>Turn on the heating pad to 37 &#176;C.</li>
	<li>Prepare the rectal temperature probe for the surgery.</li>
</ol>
2. Mouse Preparation
This step includes anesthesia, hair removal, and positioning (adult FVBN mice-age 16 - 20 weeks).
<ol>
	<li>Weigh the mice and record the weights.</li>
	<li>Anesthetize the mice.
	<ol>
		<li>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).</li>
		<li>Maintain the anesthesia with about 0.1 - 0.25 times initial doses of ketamine/xylazine throughout the surgery.&#160;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.&#160; 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.</li>
	</ol>
	</li>
	<li>Monitor the mice every 2 - 3 min by toe pinch to check for appropriate depth of anesthesia.</li>
	<li>Shave the ventral neck area of the mouse.</li>
	<li>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.</li>
	<li>Place artificial tear ointment onto the eyes to prevent drying out during surgery.</li>
</ol>
3. Surgical Details
<ol>
	<li>Make a 1 cm incision in the right ventral neck over the jugular vein.</li>
	<li>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.&#160;</li>
	<li>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.</li>
	<li>Mark a&#160;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.</li>
	<li>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.</li>
	<li>Tack the skin loosely back together around the catheter with 4-0 silk to help prevent loss of body heat and desiccation of tissues.</li>
	<li>Connect the catheter to a syringe containing heparinized saline (10 U/mL) and flush the catheter.</li>
</ol>
4. Injections
<ol>
	<li>Inject the Evans blue solution (50 &#956;L 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.</li>
	<li>2 min later, inject substance P (100 &#956;L of a 0.3 &#956;M 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.</li>
	<li>Wait 18 min after the substance P is injected. During this time, the Evans blue dye will equilibrate and circulate.</li>
	<li>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.</li>
</ol>
5. Isolation of Organs
<ol>
	<li>Cut open the chest cavity of the mouse and gravity-perfuse (from a height of about 51 cm or 20&#34;) 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.</li>
	<li>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.</li>
	<li>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).</li>
	<li>Blot the organs with tissue, cut each organ in half, and weigh each half (wet weights, in g).</li>
	<li>Dry one half of the tissue in a drying oven at 150 &#176;C, on foil, for 48 h.</li>
	<li>Place the other tissue half in a consistent volume (up to 200 &#181;L) of formamide in a microfuge tube for 48 h (and up to 72 h) to extract the Evans blue.</li>
</ol>
6. Measurement of Tissue OD
<ol>
	<li>Remove 50 &#181;L 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.</li>
	<li>Place 50 &#181;L of new, pure formamide into each of two empty wells of the 96-well plate for the blanks.</li>
	<li>Measure and record the OD620 of each well of the 96-well plate on an absorbance plate reader.&#160;620 nm is the absorbance max of Evans blue.</li>
</ol>
7. Calculation of Plasma Extravasation
<ol>
	<li>Weigh the dry tissue half which has been in the oven for 48 h.</li>
	<li>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).</li>
	<li>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).</li>
	<li>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.</li>
	<li>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.</li>
	<li>Analyze data and express as the mean &#177; SEM. Statistically compare groups by t test or one-way analysis of variance and Scheff&#233;&#39;s multiple-comparison&#160;test (lycofs01.lycoming.edu), as appropriate.</li>
</ol>

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The authors have nothing to disclose.

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...

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 desired2,3,4,5,6,7,8,9. 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 injections5,7,10, 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 injections4,9. 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&#39;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&#160;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&#160;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&#39;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.

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

an optimized evans blue protocol to assess vascular leak in the mouse

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.

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 ...

Watch this Scientific Journal Video about An Optimized Evans Blue Protocol to Assess Vascular Leak in the Mouse at JoVE.com

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

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.

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 ...

an-optimized-evans-blue-protocol-to-assess-vascular-leak-in-the-mouse

Research

JoVE Journal

Medicine

26.3K Views.  University of Colorado Denver. 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.

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

Rat Model of Blood-brain Barrier Disruption to Allow Targeted Neurovascular Therapeutics

Endothelial cells with tight junctions along with the basement membrane and astrocyte end feet surround cerebral blood vessels to form the blood-brain barrier1. The barrier selectively excludes molecules from crossing between the blood and the brain based upon their size and charge. This function can impede the delivery of therapeutics for neurological disorders. A number of chemotherapeutic drugs, for example, will not effectively cross the blood-brain barrier to reach tumor cells2. Thus, improving the delivery of drugs across the blood-brain barrier is an area of interest.
The most prevalent methods for enhancing the delivery of drugs to the brain are direct cerebral infusion and blood-brain barrier disruption3. Direct intracerebral infusion guarantees that therapies reach the brain; however, this method has a limited ability to disperse the drug4. Blood-brain barrier disruption (BBBD) allows drugs to flow directly from the circulatory system into the brain and thus more effectively reach dispersed tumor cells. Three methods of barrier disruption include osmotic barrier disruption, pharmacological barrier disruption, and focused ultrasound with microbubbles. Osmotic disruption, pioneered by Neuwelt, uses a hypertonic solution of 25% mannitol that dehydrates the cells of the blood-brain barrier causing them to shrink and disrupt their tight junctions. Barrier disruption can also be accomplished pharmacologically with vasoactive compounds such as histamine5 and bradykinin6. This method, however, is selective primarily for the brain-tumor barrier7. Additionally, RMP-7, an analog of the peptide bradykinin, was found to be inferior when compared head-to-head with osmotic BBBD with 25% mannitol8. Another method, focused ultrasound (FUS) in conjunction with microbubble ultrasound contrast agents, has also been shown to reversibly open the blood-brain barrier9. In comparison to FUS, though, 25% mannitol has a longer history of safety in human patients that makes it a proven tool for translational research10-12.
In order to accomplish BBBD, mannitol must be delivered at a high rate directly into the brain's arterial circulation. In humans, an endovascular catheter is guided to the brain where rapid, direct flow can be accomplished. This protocol models human BBBD as closely as possible. Following a cut-down to the bifurcation of the common carotid artery, a catheter is inserted retrograde into the ECA and used to deliver mannitol directly into the internal carotid artery (ICA) circulation. Propofol and N2O anesthesia are used for their ability to maximize the effectiveness of barrier disruption13. If executed properly, this procedure has the ability to safely, effectively, and reversibly open the blood-brain barrier and improve the delivery of drugs that do not ordinarily reach the brain 8,13,14.

Blood-brain barrier disruption aids the delivery of certain drugs to the brain. Mannitol delivered intra-arterially shrinks cells surrounding blood vessels in order to physically disrupt the barrier.

Endothelial cells with tight junctions along with the basement membrane and astrocyte end feet surround cerebral blood vessels to form the blood-brain ...

An in vivo Assay to Test Blood Vessel Permeability

This method is based on the intravenous injection of Evans Blue in mice as the test animal model. Evans blue is a dye that binds albumin. Under physiologic conditions the endothelium is impermeable to albumin, so Evans blue bound albumin remains restricted within blood vessels. In pathologic conditions that promote increased vascular permeability endothelial cells partially lose their close contacts and the endothelium becomes permeable to small proteins such as albumin. This condition allows for extravasation of Evans Blue in tissues. A healthy endothelium prevents extravasation of the dye in the neighboring vascularized tissues. Organs with increased permeability will show significantly increased blue coloration compared to organs with intact endothelium. The level of vascular permeability can be assessed by simple visualization or by quantitative measurement of the dye incorporated per milligram of tissue of control versus experimental animal/tissue. Two powerful aspects of this assay are its simplicity and quantitative characteristics. Evans Blue dye can be extracted from tissues by incubating a specific amount of tissue in formamide. Evans Blue absorbance maximum is at 620 nm and absorbance minimum is at 740 nm. By using a standard curve for Evans Blue, optical density measurements can be converted into milligram dye captured per milligram of tissue. Statistical analysis should be used to assess significant differences in vascular permeability.

An in vivo Assay to Test Blood Vessel Permeability

We are presenting an in vivo assay to test blood vessel permeability. This assay is based on intravenous injection of a dye and subsequent visualization of its diffusion into interstitial spaces.

This method is based on the intravenous injection of Evans Blue in mice as the test animal model. Evans blue is a dye that binds albumin. Under ...

Measuring Local Anaphylaxis in Mice

Allergic responses are the result of the activation of mast cells and basophils, and the subsequent release of vasoactive and proinflammatory mediators. Exposure to an allergen in a sensitized individual can result in clinical symptoms that vary from minor erythema to life threatening anaphylaxis. In the laboratory, various animal models have been developed to understand the mechanisms driving allergic responses. Herein, we describe a detailed method for measuring changes in vascular permeability to quantify localized allergic responses. The local anaphylaxis assay was first reported in the 1920s, and has been adapted from the technique published by Kojima et al. in 20071. In this assay, mice sensitized to OVA are challenged in the left ear with vehicle and in the right ear with OVA. This is followed by an intravenous injection of Evans Blue dye. Ten min after injecting Evans Blue, the animal is euthanized and the dye that has extravasated into the ears is extracted overnight in formamide. The absorbance of the extracted dye is then quantified with a spectrophotometer. This method reliably results in a visual and quantifiable manifestation of a local allergic response.

Allergic responses, characterized by the activation of mast cells and basophils, are driven by the cross-linking of IgE and release of proinflammatory mediators. A quantitative assessment of allergic responses can be achieved by using Evans Blue dye to monitor changes in vascular permeability after allergen challenge.

Allergic responses are the result of the activation of mast cells and basophils, and the subsequent release of vasoactive and proinflammatory mediators. ...

Immunology and Infection

Using Adeno-associated Virus as a Tool to Study Retinal Barriers in Disease

M&#252;ller cells are the principal glial cells of the retina. Their end-feet form the limits of the retina at the outer and inner limiting membranes (ILM), and in conjunction with astrocytes, pericytes and endothelial cells they establish the blood-retinal barrier (BRB). BRB limits material transport between the bloodstream and the retina while the ILM acts as a basement membrane that defines histologically the border between the retina and the vitreous cavity. Labeling M&#252;ller cells is particularly relevant to study the physical state of the retinal barriers, as these cells are an integral part of the BRB and ILM. Both BRB and ILM are frequently altered in retinal disease and are responsible for disease symptoms.
There are several well-established methods to study the integrity of the BRB, such as the Evans blue assay or fluorescein angiography. However these methods do not provide information on the extent of BRB permeability to larger molecules, in nanometer range. Furthermore, they do not provide information on the state of other retinal barriers such as the ILM. To study BRB permeability alongside retinal ILM, we used an AAV based method that provides information on permeability of BRB to larger molecules while indicating the state of the ILM and extracellular matrix proteins in disease states. Two AAV variants are useful for such study: AAV5 and ShH10. AAV5&#160;has a natural tropism for photoreceptors but it cannot get across to the outer retina when administered into the vitreous when the ILM is intact (i.e., in wild-type retinas). ShH10 has a strong tropism towards glial cells and will selectively label M&#252;ller glia in both healthy and diseased retinas. ShH10 provides more efficient gene delivery in retinas where ILM is compromised. These viral tools coupled with immunohistochemistry and blood-DNA analysis shed light onto the state of retinal barriers in disease.

To investigate the blood-retinal barrier permeability and the inner limiting membrane integrity in animal models of retinal disease, we used several adeno-associated virus (AAV) variants as tools to label retinal neurons and glia. Virus mediated reporter gene expression is then used as an indicator of retinal barrier permeability.

M&#252;ller cells are the principal glial cells of the retina. Their end-feet form the limits of the retina at the outer and inner limiting membranes ...

Inducing Ischemia-reperfusion Injury in the Mouse Ear Skin for Intravital Multiphoton Imaging of Immune Responses

Ischemia-reperfusion injury (IRI) occurs when there is transient hypoxia due to the obstruction of blood flow (ischemia) followed by a subsequent re-oxygenation of the tissues (reperfusion). In the skin, ischemia-reperfusion (IR) is the main contributing factor to the pathophysiology of pressure ulcers. While the cascade of events leading up to the inflammatory response has been well studied, the spatial and temporal responses of the different subsets of immune cells to an IR injury are not well understood. Existing models of IR using the clamping technique on the skin flank are highly invasive and unsuitable for studying immune responses to injury, while similar non-invasive magnet clamping studies in the skin flank are less-than-ideal for intravital imaging studies. In this protocol, we describe a robust model of non-invasive IR developed on mouse ear skin, where we aim to visualize in real-time the cellular response of immune cells after reperfusion via multiphoton intravital imaging (MP-IVM).

This protocol describes the induction of an ischemia-reperfusion (IR) model on mouse ear skin using magnet clamping. Using a custom-built intravital imaging model, we study in vivo inflammatory responses post-reperfusion. The rationale behind the development of this technique is to extend the understanding of how leukocytes respond to skin IR injury.

Ischemia-reperfusion injury (IRI) occurs when there is transient hypoxia due to the obstruction of blood flow (ischemia) followed by a subsequent ...

Measuring Post-Stroke Cerebral Edema, Infarct Zone and Blood-Brain Barrier Breakdown in a Single Set of Rodent Brain Samples

One of the most common causes of morbidity and mortality worldwide is ischemic stroke. Historically, an animal model used to stimulate ischemic stroke involves middle cerebral artery occlusion (MCAO). Infarct zone, brain edema and blood-brain barrier (BBB) breakdown are measured as parameters that reflect the extent of brain injury after MCAO. A significant limitation to this method is that these measurements are normally obtained in different rat brain samples, leading to ethical and financial burdens due to the large number of rats that need to be euthanized for an appropriate sample size. Here we present a method to accurately assess brain injury following MCAO by measuring infarct zone, brain edema and BBB permeability in the same set of rat brains. This novel technique provides a more efficient way to evaluate the pathophysiology of stroke.

This protocol describes a novel technique of measuring the three most important parameters of ischemic brain injury on the same set of rodent brain samples. Using only one brain sample is highly advantageous in terms of ethical and economic costs.

One of the most common causes of morbidity and mortality worldwide is ischemic stroke. Historically, an animal model used to stimulate ischemic stroke ...

Neuroscience

In Vivo Vascular Injury Readouts in Mouse Retina to Promote Reproducibility

Advancements in ophthalmic imaging tools offer an unprecedented level of access to researchers working with animal models of neurovascular injury. To properly leverage this greater translatability, there is a need to devise reproducible methods of drawing quantitative data from these images. Optical coherence tomography (OCT) imaging can resolve retinal histology at micrometer resolution and reveal functional differences in vascular blood flow. Here, we delineate noninvasive vascular readouts that we use to characterize pathological damage post vascular insult in an optimized mouse model of retinal vein occlusion (RVO). These readouts include live imaging analysis of retinal morphology, disorganization of retinal inner layers (DRIL) measure of capillary ischemia, and fluorescein angiography measures of retinal edema and vascular density. These techniques correspond directly to those used to examine patients with retinal disease in the clinic. Standardizing these methods enables direct and reproducible comparison of animal models with clinical phenotypes of ophthalmic disease, increasing the translational power of vascular injury models.

In Vivo Vascular Injury Readouts in Mouse Retina to Promote Reproducibility

Here, we present three data analysis protocols for fluorescein angiography (FA) and optical coherence tomography (OCT) images in the study of Retinal Vein Occlusion (RVO).

Advancements in ophthalmic imaging tools offer an unprecedented level of access to researchers working with animal models of neurovascular injury. To ...

A Fluorescence Assay for Evaluating the Permeability of a Cardiac Microvascular Endothelial Barrier in a Rat Model of Ischemia/reperfusion

Revascularization therapies for culprit arteries, regardless of percutaneous coronary intervention and coronary artery bypass grafting, are considered the best strategy for improving the clinical prognosis of patients with acute coronary syndrome (ACS). Nonetheless, myocardial reperfusion following effective revascularization can trigger significant cardiomyocyte death and coronary endothelial collapse, known as myocardial ischemia/reperfusion injury (MIRI). Usually, endothelial cells and their intercellular tight junctions cooperatively maintain the microvascular endothelial barrier and its relatively low permeability but fail in reperfusion areas. 
Microvascular endothelial hyperpermeability induced by ischemia/reperfusion (IR) contributes to myocardial edema, increased infiltration of pro-inflammatory cells, and aggravated intramyocardial hemorrhage, which may worsen the prognosis of ACS. The tracer used in this study-70,000 Da FITC-dextran, a branched glucose molecule labeled by fluorescein isothiocyanate (FITC)-appears too large to infiltrate the cardiac microvascular endothelium in normal conditions. However, it is capable of infiltrating a broken barrier after MIRI. Thus, the higher the endothelial permeability is, the more FITC-dextran accumulates in the extravascular intercellular space. Thus, the intensity of fluorescence from FITC can indicate the permeability of the microvascular endothelial barrier. This protocol takes advantage of FITC-dextran to evaluate the cardiac microvascular endothelial barrier functionally, which is detected by an automated quantitative pathology imaging system.

Here, we describe a method to functionally assess the endothelial barrier of cardiac microvessels after ischemia/reperfusion injury via the measurement of the mean fluorescence intensity of extravasated 70,000 Da FITC-dextran in comparison with Evans Blue.

Revascularization therapies for culprit arteries, regardless of percutaneous coronary intervention and coronary artery bypass grafting, are considered the ...

Evaluating Vascular Hyperpermeability-inducing Agents in the Skin with the Miles Assay

The primary function of the vascular endothelium in vertebrate organisms is to serve as a barrier between the blood and each tissue of the body, whereby the permeability of the endothelium to blood cells, plasma macromolecules, and water can be adapted according to the physiological need. In certain diseases, cytokines and growth factors are released that target the endothelial barrier to transiently increase vascular permeability; however, their prolonged presence may cause chronic vascular hyperpermeability and thereby tissue-damaging edema. The Miles assay is an in vivo technique that allows researchers to study vascular hyperpermeability through the proxy measurement of vascular leakage. Here, we provide a detailed protocol on how to perform this procedure in the mouse, which is the most widely used model organism to study mammalian physiology and pathology. The procedure involves the intravenous injection of Evans blue dye to label the circulating albumin followed by multiple intradermal injections of permeability-inducing agents and vehicle control solutions into opposing flanks of the mouse. Consequently, Evans blue dye gradually leaks into the dermis, where it accumulates and can be extracted for quantification as leakage induced by the permeability-inducing agent relative to the vehicle. The Miles assay can be performed in wild type or genetically modified mouse models and may be combined with drug administration to study molecular mechanisms that regulate vascular permeability and identify agents/targets capable of inducing or blocking hyperpermeability.

Here, we present a protocol to measure the vascular leakage induced by intradermal administration of permeability promoting agents into the murine skin. This technique can be used to determine the ability of molecules to promote or inhibit vascular leakage or to study the molecular mechanisms that regulate vascular permeability.

The primary function of the vascular endothelium in vertebrate organisms is to serve as a barrier between the blood and each tissue of the body, whereby ...

Biology

Fluorescent Dye-Based Visualization of Blood-Brain Barrier Breakdown in Mice

Evaluation of Blood-Brain Barrier Breakdown in a Mouse Model of Mild Traumatic Brain Injury

Fluorescent dyes are used to determine the extent of dye extravasation that occurs due to blood-brain barrier (BBB) breakdown. Labeling with these dyes is a complex process influenced by several factors, such as the concentration of dyes in the blood, permeability of brain vessels, duration of dye extravasation, and reduction in dye concentration in the tissue due to degradation and diffusion. In a mild traumatic brain injury model, exposure to blast-induced shock waves (BSWs) triggers BBB breakdown within a limited time window. To determine the precise sequence of BBB breakdown, Evans blue, and fluorescein isothiocyanate-dextran were injected intravascularly and intracardially into mice at various time points relative to BSW exposure. The distribution of dye fluorescence in brain slices was then recorded. Differences in the distribution and intensity between the two dyes revealed the spatiotemporal sequence of BBB breakdown. Immunostaining of the brain slices showed that astrocytic and microglial responses correlated with the sites of BBB breakdown. This protocol has broad potential for application in studies involving different BBB breakdown models.

A method was developed to visualize dye extravasation due to blood-brain barrier (BBB) breakdown by administering two fluorescent dyes to mice at different time points. The use of glycerol as a cryoprotectant facilitated immunohistochemistry on the same sample.

Fluorescent dyes are used to determine the extent of dye extravasation that occurs due to blood-brain barrier (BBB) breakdown. Labeling with these dyes is ...

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

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Rat Model of Blood-brain Barrier Disruption to Allow Targeted Neurovascular Therapeutics

An in vivo Assay to Test Blood Vessel Permeability

Measuring Local Anaphylaxis in Mice

Using Adeno-associated Virus as a Tool to Study Retinal Barriers in Disease

Inducing Ischemia-reperfusion Injury in the Mouse Ear Skin for Intravital Multiphoton Imaging of Immune Responses

Measuring Post-Stroke Cerebral Edema, Infarct Zone and Blood-Brain Barrier Breakdown in a Single Set of Rodent Brain Samples

In Vivo Vascular Injury Readouts in Mouse Retina to Promote Reproducibility

A Fluorescence Assay for Evaluating the Permeability of a Cardiac Microvascular Endothelial Barrier in a Rat Model of Ischemia/reperfusion

Evaluating Vascular Hyperpermeability-inducing Agents in the Skin with the Miles Assay

Evaluation of Blood-Brain Barrier Breakdown in a Mouse Model of Mild Traumatic Brain Injury