Microperfusion Technique to Investigate Regulation of Microvessel Permeability in Rat Mesentery

Summary

The modified Landis technique enables paired measurement of the hydraulic conductivity of individual microvessels in the mesentery of normal and genetically modified rats under control and test conditions using microperfusion techniques. It provides a convenient method to evaluate mechanisms that regulate microvessel permeability and transvascular exchange under physiological conditions.

Abstract

Experiments to measure the permeability properties of individually perfused microvessels provide a bridge between investigation of molecular and cellular mechanisms regulating vascular permeability in cultured endothelial cell monolayers and the functional exchange properties of whole microvascular beds. A method to cannulate and perfuse venular microvessels of rat mesentery and measure the hydraulic conductivity of the microvessel wall is described. The main equipment needed includes an intravital microscope with a large modified stage that supports micromanipulators to position three different microtools: (1) a beveled glass micropipette to cannulate and perfuse the microvessel; (2) a glass micro-occluder to transiently block perfusion and enable measurement of transvascular water flow movement at a measured hydrostatic pressure, and (3) a blunt glass rod to stabilize the mesenteric tissue at the site of cannulation. The modified Landis micro-occlusion technique uses red cells suspended in the artificial perfusate as markers of transvascular fluid movement, and also enables repeated measurements of these flows as experimental conditions are changed and hydrostatic and colloid osmotic pressure difference across the microvessels are carefully controlled. Measurements of hydraulic conductivity first using a control perfusate, then after re-cannulation of the same microvessel with the test perfusates enable paired comparisons of the microvessel response under these well-controlled conditions. Attempts to extend the method to microvessels in the mesentery of mice with genetic modifications expected to modify vascular permeability were severely limited because of the absence of long straight and unbranched microvessels in the mouse mesentery, but the recent availability of the rats with similar genetic modifications using the CRISPR/Cas9 technology is expected to open new areas of investigation where the methods described herein can be applied.

Introduction

Microperfusion in the vasculature entails establishing controlled flow of an artificial perfusate of known composition via a micropipette in a blood vessel usually less than 40 µm in diameter. The perfused vessel remains within its normal tissue environment and is perfused with the animal’s blood up to the time of cannulation. When used in conjunction with a range of video imaging or fluorometric techniques, in situ microperfusion enables measurement of water and solute flows across the walls of microvessels under conditions where the driving forces for these flows are known and the permeability properties of the vascular wall can be directly evaluated. Further, by controlling the composition of the fluid surrounding the microvessel in the tissue (perfusate and superfusate), the regulation of microvessel permeability and exchange can be investigated by enabling the endothelial cells forming the microvessel wall to be exposed to a variety of experimental conditions (agonists, modified perfusion conditions, fluorescent indicators to measure intracellular composition and signaling) for precisely measured periods of time (sec to hr). In addition, ultrastructural or cytochemical evaluations of key cellular molecular structures regulating the barrier can be investigated in the same microvessels in which permeability is directly measured. The approach thereby forms a bridge between investigation of the cellular and molecular mechanisms to modify endothelial barrier function in cultured endothelial cell monolayers and investigation in intact microvessels. See the following reviews for further evaluation^1-6.

A limitation of microperfusion is that it can be used only in microvascular beds that are thin, transparent and have sufficient structural integrity to enable cannulation with a glass micropipette. While early investigations used frog microvessels in mesentery and thin cutaneous pectoris muscle^7,8, by far the most commonly used preparation in mammalian models is the rat mesentery^9-15. Most investigations have focused on acute changes in vascular permeability studied over periods of 1-4 hr, but more recent investigations have been extended to measurements on individual vessels 24-72 hr after an initial perfusion^12,16. The recently developed CRISPR technology, which promises to make more genetically modified rat models available for studying vascular permeability regulation¹⁷ should enable the methods described in this communication to be applied in venular microvessels of the mesentery in these important new rat models.

The method requires an inverted microscope equipped with a custom built microscope stage large enough to hold both the animal preparation and at least three micromanipulators used to position microtools close to the perfused vessel and to align a perfusion micropipette with the vessel lumen. For example a custom platform for an x-y microscope stage (about 90 × 60 cm) can be fabricated from a 1 cm thick steel plate with a rust-proof coating. The stage is attached to an engineering index table or two dove-tail slides mounted at right angles and supported on Teflon pillars or ball transfers for movement in the horizontal plane. A typical rig (see Figure 2) has much in common with the microscope and micropositioning equipment used for a range of intravital microcirculation experiments such as those to measure single vessel blood flow and hematocrit, local oxygen delivery by blood perfused microvessels, regulation of vascular smooth muscle tone, and the local microvascular accumulation of fluorescent tracers injected into the whole circulation.^18-26

The fundamental aspect of the technique is the measurement of volume flow (J_v) across a defined surface area (S) of the microvessel wall. To accomplish this via the modified Landis technique described herein a simple inverted microscope is adequate. A small video camera is mounted on the image port and the video signal, with an added time base, is displayed on a video monitor and recorded either in digital form on a computer or as a digital or analog signal on a video recorder. Once the microvessel is cannulated the portion of the microvessel visible to the camera can be changed by moving the stage and manipulators as a unit without disrupting the cannulation.

Measurement of transvascular flows may also be combined with more detailed investigations using a sophisticated fluorescence microscope with appropriate filters such as rigs used for measurements of solute permeability, fluorescent ratio monitoring of cytoplasmic calcium or other cellular mechanisms, and confocal imaging ^6,12,13,27. A key advantage of all microperfusion approaches is the ability to make repeated measures, on the same vessel, under controlled change of driving force such as hydrostatic and oncotic pressures, or induced change in vessel responses to inflammatory conditions. The most common design is a paired comparison of measured hydraulic conductivity (L_p) on the same vessel with the vessel first perfused via a micropipette filled with a control perfusate and the red cell suspension to establish a baseline permeability state, then with a second pipette with the test agent added to the perfusate. Multiple cannulations are possible with the cycle repeated after reperfusion with the control pipette.

The present protocol demonstrates the cannulation and microperfusion of a venular vessel in rat mesentery to record water fluxes across the microvessel wall and measure the L_p of the vessel wall, a useful index of the permeability of the common pathway for water and solutes across the intact endothelial barrier. The procedure is called the modified Landis technique because the original Landis principle of using the relative movement of red cells as a measure of transvascular fluid exchange after perfusion is blocked is preserved²⁸, but the range of experimental conditions (e.g., the hydrostatic and albumin oncotic pressure differences across the microvessel wall) available after microperfusion is far greater than in uncannulated blood perfused microvessels^8,29.

Protocol

Ethics Statement: All procedures were reviewed and approved by the Institutional Animal Care and Use Committee.

1. Preliminary Fabrication of Micropipettes, Restrainers, and Blockers

1. Pull several clean borosilicate glass capillary tubes in half using an electronic puller adjusted so that, when pulled, the stretched portion of the tube is about 1 cm in length and the two halves are somewhat symmetric. Ensure that the taper is consistent with the dimensions in Figure 1. Use both halves for micropipettes, restrainers and blockers.
   1. Bevel the micropipettes using an air driven grinding wheel³⁰ with 0.5 µm abrasive paper bathed with water. Set the angle of the micropipette holder so that it is about 30 degrees from horizontal.
   2. Thoroughly wash and dry the micropipettes by using suction to pull clean acetone through the tip and up the shaft. Inspect the micropipettes (Figure 1A) using a small upright microscope with 4× and 40× objectives fitted with an alligator clip micropipette holder and an eyepiece reticle.
   3. Select micropipettes with tip opening about 50 µm long with a bevel whose length/width ratio is between 3.1 and 3.5 and with a sharply tapered point which helps penetrate the tough collagen fibers in the mesentery.
   4. Store 15-20 micropipettes of slightly varying sizes in a dust-free box.
2. Make restrainers using pulled capillary tubes prepared in step 1.1). Hold a pulled glass capillary tube briefly near a microflame to form a blunt end (Figure 1B). Store several in a dust-free box.
3. Make microoccluders using pulled capillary tubes prepared in step 1.1). Hold a pulled capillary tube under a microflame and gently bend (using a tubing adapter, e.g., 23G) the fine tip (about 4 mm from tip) to make an angle of close to 120 degrees to the shaft (Figure 1C). Make and store several.

2. Animal Preparation and Surgery

1. Anesthetize male (350-450 g) or female Sprague-Dawley rats (200-300 g) with pentobarbital sodium (80-100 mg/kg, Sigma-Aldrich, P3761) via subcutaneous injection; protect the mesentery by not using intraperitoneal injection.
2. Throughout surgical procedures and experimental protocols, maintain anesthesia by supplemental injections (30 mg/kg) as needed. Determine depth of anesthesia by toe pinch or corneal reflex. After completion of the experimental procedure, euthanize the rat via pentobarbital overdose or intra-cardiac injection of saturated potassium chloride under anesthesia.

3. Prepare Solutions and Erythrocytes for use as Flow Markers

1. Prepare mammalian Ringer's solution for superfusate and control perfusate solution (typically mammalian Ringer's solution containing 10 mg/ml fatty acid free bovine serum albumin, BSA).
   1. Filter perfusate through 0.2 µm syringe filters to remove tiny particles that might block the micropipette tip; filter into small clean container.
2. Prepare erythrocytes by drawing about 0.2 ml blood from the tail vein of the anesthetized rat into a 20G needle on a small syringe containing 0.05 ml heparin (1,000 U/ml) and transfer to a 15 ml centrifuge tube containing about 14 mL mammalian Ringer's solution solution.
   1. Centrifuge to gently pack red cells (about max 200 × g for 7 min). Draw off the supernatant and re-suspend the red cells in Ringer's solution.
   2. After centrifuging and removing supernatant three times leave the packed red cells in the bottom of the tube for later use. Note that this procedure reduces platelets in the final perfusates to less than 0.1% of normal levels³¹.

4. Arrange the Tissue on the Animal Tray

1. Shave the abdomen of the anesthetized rat from below navel to xiphoid process. Ensure that the shaved area extends around the right side (for rat placed on right side) so that hair does not form a wick to draw the superfusate out of the tissue well. Remove cut hairs with a wetted gauze pad or tissue.
   1. Hold the skin with a blunt serrated forceps and make a center-line incision (2-3 cm long) through the skin using a sturdy scissors. Using a fine (iris) scissors make a similar incision through the linea alba, lifting the abdominal wall with serrated forceps to avoid damaging the gut.
   2. With the animal on its side on the animal tray, gently pull out the gut from the abdominal cavity using cotton-tipped applicators (wood stick) soaked in Ringer's solution and blunt serrated forceps. Arrange the gut in the tissue well so that the mesentery is draped over the pillar for viewing through the microscope taking care to avoid touching the mesentery (potentially damaging microvessels) with either forceps or cotton.
      Note: The animal tray (Figure 2A) can be constructed from Plexiglas and requires a tissue well (about 3 mm deep), an optically clear pillar (e.g., polished quartz about 2 cm dia. and 5 mm high), and a warming pad adjusted to maintain the animal’s temperature. The thickness of the pillar must not exceed the working distance of the objective.
2. Position the animal tray on the microscope stage so that the mesentery is visible through the eyepieces.
3. Position a gravity-fed drip line to continuously superfuse the mesentery with mammalian Ringer's solution maintained at 35–37 °C. Use gauze pads to hold the gut in place, help retain moisture, and wick excess Ringer's solution off the surface. Adjust the flow and aspirate the effluent to maintain a consistent superfusate thickness.
4. Identify target venular microvessels, which are unbranched segments downstream of convergent flow, one or two branches distal to true capillaries. Find an unbranched vessel (ideally, 600 to 1,000 µm in length) having brisk blood flow and free of white cells sticking on the vessel wall.
5. Position the test microvessel in the center of the microscope field and move the restrainer into position near the chosen cannulation site.

5. Fill Micropipette and Mount in Holder

1. Just before cannulating, suspend the red cells in the perfusate. Add 7 µl of packed red cells to 0.5 ml perfusate in a clean borosilicate test tube. Note: Hematocrits of 1-3% can be used, but consistent hematocrit will lead to consistent perfusate concentrations of any substances released from the red cells.
2. Fit a 1 ml syringe with a 23G tubing adapter and PE 50 tubing (5-15 cm length). Draw the perfusate/red cell mixture into the syringe. Invert syringe to mix and expel all bubbles from tubing and adapter. For increased efficiency, fit tubing to several syringes before the experiment starts and keep them in a dust free box or plastic bag.
3. Fill the micropipette by advancing the tubing into the large end of the micropipette until it abuts the tapered region. Apply a gentle quick push on the syringe plunger to fill the micropipette tip. Note: A tiny stream or droplet should be visible at the tip to evidence complete filling.
4. Withdraw the tubing while gently pushing on the plunger to fill the wider part of the micropipette shaft. Remove small bubbles in the wide shaft by carefully flicking the micropipette shaft. Note: A similar procedure has been illustrated³².
5. Place the micropipette into a pipette holder with a side port which allows a continuous fluid connection to a water manometer. Ensure that the holder, which is attached to a hydraulic drive used to control very fine advancement of the micropipette, is at a slight angle (15-25°) to the horizontal so that the edge of the micropipette taper does not hit the gut or tray.
6. Mount the hydraulic drive on a moveable micromanipulator, which has x, y, and z drives to enable close alignment to the test vessel (Figure 2).
7. Adjust the hydrostatic pressure applied to the fluid in the micropipette using a syringe or water-filled rubber bulb to change the height of fluid in the water column of the manometer to about 40 cmH₂O above the mesentery.
8. Cannulate the microvessel as soon as possible after filling the pipette; red cells settle to the bottom of the pipette, so that after about 40 min very few red cells will flow into the vessel.

6. Microcannulation and Microvascular Pressure Measurement

1. Before positioning the filled micropipette under the microscope, gently press the restrainer onto the tissue near the microvessel applying sufficient force to grip the tissue. Draw the restrainer back, slightly stretching the tissue in line with the microvessel so that the stresses in the mesenteric collagen fibers are aligned with the vessel.
   1. Align the micropipette with the vessel and lower it into view through the eyepieces. Place the tip just upstream of the chosen cannulation site and lower it onto the tissue so that it partially obstructs flow within the vessel, but does not occlude it.
2. Cannulate the vessel by driving the pipette tip slowly into the vessel lumen using the hydraulic drive. Be careful not to push through the lower side of the vessel.
   Note: As the micropipette enters the lumen, the perfusate rapidly washes the blood in the vessel out of the lumen and perfusate freely flows into the animal’s circulation distal to the test microvessel, displacing some of the normal blood perfusion in the downstream vessels.
3. Lower the perfusion pressure by adjusting the manometer until the blood (as indicated by the animal’s red cells) is drawn back into the perfused segment; this determines the balance pressure where the red cells gently oscillate in the vessel lumen and measures the hydrostatic microvessel pressure (P_c) at the distal end of the microvessel segment. Maintain vessel perfusion at all pressures above this balance pressure.

7. Microocclusion and Collection of Data

1. Optional step: Before using the micro-occluder to block the vessel, press it into the tissue in a non-used area of mesentery far from any vessel so that the tip accumulates a fine pad of tissue that protects the experimental vessel during repeated micro-occlusions.
2. Place the blocker above the perfused vessel near the lower edge of the microscope field.
3. Record the following with video and audio.
   1. Verbally record the location of the block site and the lower screen edge as seen on the eyepiece reticle.
   2. With the tip of the blocker placed just above the microvessel, use the fine z control of the micromanipulator to gently lower the blocker until the flow in the vessel is occluded. Note the manometer pressure (P_c) on audio. Apply occlusion typically for 3-10 sec, and then release, restoring free perfusion. Move the block site up the vessel toward the pipette tip in 5-10 µm steps based on time (>5 min after initial block at a site), number of occlusions (3-5), to prevent vessel wall damage at the block site.

8. Analysis of Data and Measurement of L_p (Water Permeability)

1. During video playback, determine the initial distance (ℓ₀) from a marker red cell to the occlusion site by use of an image of a stage micrometer and the known positions on the images. Determine the initial velocity (dℓ/dt) of the marker cell from its position in several frames during the 3-10 sec of recorded images. Determine the vessel radius (r) from the images taken during occlusion (Figure 3).
2. Calculate J_v/S for each occlusion as (dℓ/dt)×(1/ℓ₀)× (r/2). Calculate L_p at a constant pressure as (J_v/S) divided by the driving pressure (microvessel pressure – effective colloid osmotic pressure; see Discussion).

Results

Figure 4 shows the results from measuring the time course of changes in L_p in a rat venular microvessel cannulated successively with four perfusates.³³ The magnitude of L_p calculated at a constant pressure was used as a measure of changes in microvessel wall permeability, first in the control state with a perfusate containing 1% bovine serum albumin then when the vessel was exposed to the inflammatory agent bradykinin (Bk) using a second micropipette containing 10 nM Bk. ...

Discussion

Details of L_p calculations. Although transvascular fluid movement occurs while the vessel is freely perfused, such exchange is far too small to be measured during free perfusion because it is typically less than 0.01% of the vessel perfusion rate. However, when perfusion is transiently stopped by occluding the microvessel, transvascular flow (i.e., filtration) is measured from the movement of marker red cells in the lumen as the column of fluid between a marker red cell and ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
MICROSCOPE, TABLE AND STAGE
inverted microscope (metallurgical type) with trinocular head for video: example	Olympus	CK-40	try to place eyepieces higher relative to stage--you have to look through eyepieces while reaching around to top of stage over intervening micromanipulators
inverted microscope (metallurgical type) with trinocular head for video: example	Leica	DMIL	try to place eyepieces higher relative to stage--you have to look through eyepieces while reaching around to top of stage over intervening micromanipulators
narrow diameter, long working distance objective: example	Nikon	Nikon E Plan 10×/0.25 LWD
stage platform--1/2 inch or 1 cm sheet steel	welding shop		this should be heavy to reduce vibration
Unislide x-y table: dove tail slides	Velmex	AXY4006W1
VIDEO
CCD video camera: example	Pulnix	TM-7CN (no longer available)	no color needed
video capture system with audio--generic
video playback system (completely still frame, single frame motion)
small microphone
MICROMANIPULATORS, HOLDERS
micromanipulator, XYZ (3)	Prior/Stoelting (no longer available)		look for fine Z, and larger range of travel in coarse drives for ease of positioning
hydraulic probe drive, one way	FHC	50-12-1C	need to buy either manual drive or electronic drive
manual drum drive	FHC	50-12-9-02
or hydraulic drive, 3 way	Siskiyou Corporation	MX610 (1-way) or MX630 (3-way)	great for short arms, water filled and must be sent back for refill ~every 2 years
connectors/rods/holders	Siskiyou Corporation	MXC-2.5, MXB etc.
pin vise	Starrett	162C	to hold restrainer
pipette holder	World Prescision Instruments	MPH3
water manometer ~120 cm
MICROSCOPE TRAY
clear Plexiglas for microscope tray for animal
3/4 inch polished quartz disc ~1/4 inch tall	Quartz Scientific Inc.	custom	(or polished plexiglass, glass); make sure the height is less than working distance of objective
Plexiglas glue (Weld-on 4: CAUTION CARCINOGEN)
medical adhesive for tissue well	NuSil	MED-1037
All-purpose silicone rubber heat mat, 5" L x 2" W	Cole Parmer	EW-03125-20	heater for microscope tray--needs cord and controller--240V version available
Power Cord Adapter for Kapton Heaters and Kits, 6 ft, 120 VAC	Cole Parmer	EW-03122-75
STACO 3PN1010B Variable-Voltage Controller, 10 A; 120 V In, 0-140 V Out	Cole Parmer	EW-01575-00
PIPET MANUFACTURE
vertical pipette puller	Sutter Instrument Company	P-30 with nichrome filament
1.5 mm OD thin wall capillary tubing	Sutter Instrument Company	B150-110-10
pipette grinder air stone and dissection microscope--see reference in text			or purchase a package from Sutter Instruments or World Precision Instruments
RX Honing Machine, System II	RX Honing Machine Corporation	MAC-10700 Rx System II Machine	alternative for air stone, use with a dissecting microscope mounted at an angle
with ceramic sharpening disc	RX Honing Machine Corporation		use "as is" or attach lapping film
lapping film sheets, 0.3 or 0.5 um	3M	part no. 051144 80827	268X Imperial lapping film sheets with adhesive back--can be purchased from Amazon