Quantification of Diabetes-induced Adherent Leukocytes in Retinal Vasculature

Summary

We demonstrate a method to label the walls of the retinal vasculature and adherent leukocytes. These adherent leukocytes can then be counted under a fluorescence microscope as a parameter of inflammation or the response of that inflammation to therapies.

Abstract

Leukostasis refers to the attachment of leukocytes to the luminal wall of the vasculature. This interaction of leukocytes with the wall of blood vessels is characteristic of inflammation and has been causally linked to capillary occlusion in a variety of tissues and diseases, including diabetic retinopathy.

Leukostasis has been reported for years as a life-threatening complication of hyperleukocytosis and can only be diagnosed clinically. Given the importance of the phenomenon, intensive research has been done to understand the potential mechanism(s) that lead to its manifestation; however, there is no gold-standard technique in laboratory settings to visualize and quantify the severity of the event.

In the method summarized below, the vasculature is initially perfused with a buffer to remove blood, and then, concanavalin A is perfused into the vasculature where it binds to all exposed cell walls and causes especially bright staining of leukocytes. If the perfusion to remove all unbound blood cells was successful, the remaining fluorescently labeled leukocytes are bound to the vasculature, and they can be manually quantified using any available fluorescence microscope.

Introduction

Leukocytes (white blood cells, WBCs) play an important role in the optimal function of the vasculature such as maintenance of the blood fluidity and regulation of thrombus resolution¹. They also play a key role in some pathological conditions, such as adhering to the luminal wall of the vasculature for prolonged periods of time leading to vessel obstruction, at least temporarily, a phenomenon known as leukostasis^2,3.

Diabetic retinopathy is one of the most common complications of long-term diabetes and one of the leading causes of visual impairment and blindness in the US and worldwide for individuals 20-75 years of age⁴. Slow and progressive degeneration of the retinal vasculature is a clinically meaningful component of the early stages of the disease, which in some patients leads to retinal ischemia with the resulting retinal neovascularization^5,6. Cumulative evidence indicates that inflammation plays an important role in the development of the retinopathy⁷, and leukostasis is considered a subclinical intravascular inflammatory response. Leukostasis occurs in the early stages of diabetes, well before any detectable clinical manifestations have developed^8,9,10. The repeated plugging of the retinal vessels by adherent leukocytes over months to years (chronic leukostasis) in diabetes might contribute to the vascular occlusion and degeneration of the capillaries^11,12,13. The severity of this leukostasis is of pathologic significance and can be used to monitor the severity of the disease process or to evaluate the efficacy of a therapy in research settings.

To further study the specific effects of the hyperglycemic microenvironment on leukostasis, in vitro models have been designed. Isolated retinal microvascular endothelial cells can be grown and arranged either in 2- or 3-D cultures models (microvasculature-on-a-chip¹⁴) to replicate the vascular endothelium (the cell monolayer that paves the lumen of the vessels). However, the interexperimental variation of these models limits their use. The study of leukostasis in human retinal vasculature in vivo is still limited, and therefore, most of the current knowledge on retinal leukostasis is derived from animal models of diabetic retinopathy^13,15.

The aim of this report is to describe a standard protocol based on methods described elsewhere¹⁶ for the quantification of attached leukocytes to the retinal vasculature as a parameter of leukostasis. This assay can be used to study other vascular diseases that also present leukostasis, such as malignancies^3,17,18,19 and some infectious and allergic conditions²⁰. This protocol can be implemented in any basic research laboratory without the need of specialized equipment. In the method summarized below, the vasculature is initially perfused with buffer to remove blood, and then, concanavalin A is perfused into the vasculature where it binds to all exposed cell walls and causes especially bright staining of leukocytes^21,22,23. If the perfusion to remove all unbound blood cells is successful, the remaining fluorescently labeled leukocytes that are bound to the vasculature can be manually quantified using any fluorescence microscope on hand.

Protocol

The protocol has been reviewed and approved by the Institutional Animal Care and Use Committee (IACUC) at the University of California Irvine and conforms to governmental regulations regarding the care and use of laboratory animals. There are no stop points in this protocol. The average time per mouse is 30 min.

1. Preparing the perfusion stage

1. Warm up the 0.9% saline bag and concanavalin A solution in a 37 ^oC water bath for 20-30 min before use.
   NOTE: Protect concanavalin A from light exposure (cover with foil).
2. Set up a tray to contain blood and liquids from dripping on the surface where the procedure will take place. On top of the tray, place a heating pad covered with an absorbent bench underpad or any absorbent material.
   NOTE: The goal is to avoid the mouse's body losing heat during the procedure, because cooling makes more difficult to remove blood during perfusion.

2. Setting up the pressure infuser

1. Connect in series the bag of 0.9% saline, the I.V. catheter set, a 4-way valve stopcock, and the gavage needle.
2. Insert the 0.9% saline bag between the netting and the air bladder of the pressure infuser. Hang the saline bag on the hook located on the back of the air bladder. Use the I.V. pole loop to hang the pressure infuser in the I.V. pole.
3. Purge the lines and ports of all air bubbles by letting the system open (run) for a couple of minutes and set the flow rate to 18-20 mL/min²⁴. To inflate the pressure infuser air bladder, turn the stopcock handle to point toward the open stopcock vent, then pump the inflation bulb until the pressure gauge indicates the desired pressure. Re-adjust the pressure before perfusing each mouse. To deflate, turn the stopcock handle straight down towards the inflation bulb.
   NOTE: If the 0.9% saline bag is new, usually 150 mmHg pressure delivers the desired flow rate; however, the pressure should be adjusted empirically due to variations in pressure infuser brands and over the period of use of the 0.9% saline bag.
4. Attach a 10 mL syringe filled with warmed concanavalin A solution to the 4-way valve.
   NOTE: Protect the syringe from light exposure (cover with foil).

3. Anesthesia

1. Deliver anesthesia by intraperitoneal (I.P.) injection of Ketamine:Xylazine; the most widely used dose for mouse surgery/procedure is 100:10 mg/kg body weight²⁵. Assess anesthesia by pedal reflex (firm toe pinch).
   NOTE: This dose provides an onset of 4-6 min with a 45-60 min duration of surgical anesthesia. The anesthetic cocktail can be stored at room temperature for a maximum of 2 weeks.

4. Transcardial perfusion and staining with concanavalin A

1. Place the mouse on the perfusion stage in the supine position to allow for exposure of the thoracic and abdominal cavity.
2. Visually identify the xiphoid process, and with the hemostat in the dominant hand, pin the skin and lock it. Once the hemostat is secured, transfer it to the nondominant hand and lift the skin.
3. Use scissors in the dominant hand and cut, in a 90° angle to the spine, a patch of skin to reveal the outer abdominal wall.
4. With the xiphoid process and the rib cage now visible, dissect through the abdominal wall bilaterally, taking care to avoid cutting any organs or major vessels.
5. With the diaphragm now visible, visualize the heart ventricles and lungs through the diaphragm. Using the tip of the scissors, cut through the diaphragm in one of the flanks, close to the spine, taking care to avoid cutting any organs or major vessels.
   NOTE: This "hole" in the diaphragm will equilibrate the negative intrathoracic pressure with the atmospheric pressure, and a pneumothorax will occur collapsing the lungs and retracting the heart, facilitating the dissection of the diaphragm without damaging the lungs or the heart.
6. Continue dissecting through the ribs and parallel to the lungs to create a chest "flap". Release the hemostat and cut the xiphoid process in the sagittal plane. Gently wide open the xiphoid process manually. Observe the four chambers of the heart.
7. With the nondominant hand and using forceps, grasp the heart near its apex. With the dominant hand, hold the gavage needle (attached to the I.V. catheter) and puncture the apex of the heart. To avoid full perforation of the left ventricle or reaching the pulmonary vasculature and then poor perfusion of the systemic vasculature, check the placement of the end of the ball tip of the gavage needle, which should be at the edge of the site of puncture slightly protruding from the heart. Clamp the gavage needle in place using curved mosquito forceps or simply hold it by hand while manipulating the I.V. stopcock.
8. Open the stopcock to the 0.9% saline and almost simultaneously, cut open the right ventricle with scissors; perfuse for 2-3 min. During the perfusion time, gently move the needle from side to side and up and down to reduce kinking of the vasculature and increase blood exit from the heart.
9. After perfusing with saline, turn the stopcock handle to shut off flow from the saline and allow flow from the syringe to the gavage needle. Perfuse by hand with the concanavalin A solution at a steady state rate. Ensure the 10 mL of concanavalin A solution is dispensed in 30-35 s.
10. After perfusing with concanavalin A, turn the valve to shut off flow from the syringe and allow flow from the 0.9% saline to the gavage needle again. Perfuse with the 0.9% saline solution for an additional 2-3 min. Remove the gavage needle from the heart.
    NOTE: The concanavalin A suggested in this protocol is conjugated to fluorescein (green); however, concanavalin A attached to other fluorochromes is also available.

5. Enucleation and isolation of fresh retina

1. Turn the mouse on its side and, using the nondominant hand, place the index finger and thumb on the superior and inferior eyelids, respectively. Gently retract the eyelids and skin with fingers and proptose the eye, making it partially bulge out of the socket.
2. While the eye is proptosed, use curved scissors in the dominant hand and scoop under the eye in a 45° angle. Cut the muscular attachment and optic nerve. Using the same scissors as a spatula, transfer the eye to a small container or directly to the stage of the dissecting microscope.
   NOTE: Be careful not to cut off the back of the eye and avoid pulling the eye during this step.
3. Place the eye on a dental wax to open the globe. Under the dissecting microscope and using the nondominant hand, hold the scleral fold or the muscle remnants still attached externally to the posterior eye with micro-forceps, and orient the eye so that the cornea faces to a side.
   NOTE: To prevent the eye from moving/sliding while opening the globe, a piece of wet lint-free tissue can be placed on top of the dental wax.
4. With one of the sharp corners of a Teflon-coated razor blade, make an incision 1-2 mm behind and parallel to the limbus (cornea-sclera junction). Hold the scleral fold or muscle with the micro-forceps, and draw the blade across the limbus with minimal downward force. Continue to cut with the razor to totally separate the anterior segment (cornea, iris, lens, and vitreous) from the posterior segment (eye cup).
   NOTE: Do not saw back and forth.
5. Transfer the bisected eye cup to a small Petri dish with PBS.
   NOTE: Avoid contact of the retina with the tissue paper (note in step 5.3) since it will tightly stick to the paper and become essentially impossible to recover.
6. Grab a scleral fold or the remaining muscle on the outside of the sclera with micro-forceps. Completely detach the retina from the sclera by breaking all the connections at the limbus around the perimeter of the eye cup using a micro-spatula. Scoop the retina out from the sclera with the micro-spatula. If the retina is still attached to the sclera by the optic nerve, slip the micro-scissors between the retina and sclera to cut the optic nerve.
7. Remove any remnants of vitreous and ciliary muscle in the periphery of the retina. Immediately transfer the isolated retina to a slide with some PBS.
   NOTE: Any other retina isolation technique can be used depending on the researcher's preference.

6. Flat mounting of the retina

1. Lay out the unfixed retina on a slide with a small amount of PBS. Using the micro-spatula, gently orient the retina with the vitreous side up. If the retina is folded inward, use micro-forceps to hold the edges of the retina while the retina is unfolded using the micro-spatula.
2. Make 4-5 radial cuts into the retina so that it lies flat (cloverleaf pattern).
3. Using a lint-free tissue, dry up the excess of PBS away from the retina.
   NOTE: Do not touch the retina with the tissue; otherwise, the sample will be lost. Coverslipping is desirable to keep the retina flat.

7. Microscopy

NOTE: Any fluorescence microscope with a GFP/FITC (480/530 nm) channel can be used for this step. For this work, we used the referenced microscope with 488 channel and associated software for image acquisition.

1. Observe the recently flat-mounted retina under the microscope at 100x magnification (10x objective) and count fluorescently labeled leukocytes (manually) by methodically scanning the entire tissue (right to left or top to bottom).
   NOTE: Leukocytes are single fluorescent dots that can display a round or oval shape. They are 12-15 µm in diameter and do not protrude from the retinal capillaries (the structure is completely constrained by the lumen of the vessel).
2. Acquire representative images with the desired magnification and perform postprocessing of the images with the software of choice (e.g., ImageJ [Fiji]).
3. Express the count as leukocytes per retina. Graph the data by mean ± standard deviation.

Results

A well-executed perfusion and staining protocol will show the complete retinal vasculature delineated with concanavalin A (Figure 1). Poor perfusion of the mouse prevents labeling of the entire vascular tree and subsequent analysis of the leukocytes adherent to the lumen (Figure 2), whereas excessive pressure from a rapid squeeze of a syringe (less than 30-35 s) can cause vascular permeability and bursting of the blood vessels (Figure 3

Discussion

Leukostasis in humans refers to symptoms and clinical findings associated with hyperleukocytosis (total leukocytes (WBCs) count >100,000/µL) and is a medical emergency²⁰. The mechanism(s) that lead to leukostasis are under intensive research. To date, the study of leukostasis in humans in vivo is not yet possible and researchers need to rely on animal models to understand this process. Different diseases present leukostasis and having a detailed protocol to visualize the phenomen...

Materials

Name	Company	Catalog Number	Comments
10 mL syringe
4-way stopcock Luer lock I.V. line valve	Baxter	2C6204
Concanavalin A solution	Vector	FL-1001	Prepare in PBS 1 mg/mL
Dissecting tools set			Includes hemostats, scissors and forceps
FIJI			Software for image processing
Fluorescence microscope	Nikon	Eclipse Ni
Forceps, Dumont #5, Biological grade tip	Electron Microscopy Sciences (EMS)	72700-D
Gavage Needle 1.25 mm OD barrel tip x 30 mm	Fine Science	18060-20
Halstead Mosquito Forceps	Fisher Scientific	13-812-10
I.V. Catheter set with regulating clamp 70 inches	Baxter	2C5417s
I.V. Pole
Lint free tissue			Kimpwipes is an option
Micro dissecting spring scissors, Vannas, 3 mm straight	ROBOZ	RS-5620
Micro spatula	Fine Science Tools (FST)	10091-12
Nikon		NIS-Elements (AR 5.30.03 64-bit)	Software for image acquisition
Petri dish (100 mmx15 mm)	Corning	351029
Phosphate buffered saline (PBS)
Pink dental wax	Electron Microscopy Sciences (EMS)	72670
Pressure infuser	Infusurge	4010
Razor blades, GEM single edge stainless steel, Teflon coated	Electron Microscopy Sciences (EMS)	71970
Saline 0.9%, veterinary grade, 1000 mL	Baxter	04925-04-10
Small dissecting scissors, curved blunt end 22 mm	ROBOZ	RS 5983